Cloﬁbrate causes an upregulation of PPAR ... - uni-halle.de

Clofibrate caus es an upregulation of PPAR-� target genes but does not alter

expres s ion of S REB P target genes in liver and adipos e tis s ue of pigs

Sebastian Luci, Beatrice Giemsa, Holger Kluge, and Klaus Eder

Institut fur Agrar- und Ernahrungswissenschaften, Martin-Luther-Universitat Halle-Wittenberg, Halle (Saale), Germany

S ubmitted 25 Augus t 2006; accepted in final form 7 March 2007

Luci S, Giemsa B, Kluge H, Eder K. Clofibrate caus es anupregulation of PPAR-� target genes but does not alter expres s ion ofS REB P target genes in liver and adipos e tis s ue of pigs . Am J PhysiolRegul Integr Comp Physiol 29 3 : R000 –R000, 2007 . Firs t publis hedMarch 1 5 , 2007 ; doi: 1 0. 1 1 5 2/ajpregu. 00603 . 2006. —This s tudy inves -tigated the effect of clofibrate treatment on expres s ion of target genesof peroxis ome proliferator- activated receptor ( PPAR) -� and variousgenes of the lipid metabolis m in liver and adipos e tis s ue of pigs . Anexperiment with 1 8 pigs was performed in which pigs were fed eithera control diet or the s ame diet s upplemented with 5 g clofibrate/kg for28 days . Pigs treated with clofibrate had heavier livers , moderatelyincreas ed mRNA concentrations of various PPAR-� target genes inliver and adipos e tis s ue, a higher concentration of 3 - hydroxybutyrate,and markedly lower concentrations of triglycerides and choles terol inplas ma and lipoproteins than control pigs (P � 0. 05 ) . mRNA con-centrations of s terol regulatory element- binding proteins ( S REB P) - 1and - 2, ins ulin- induced genes (Insig) - 1 and Insig-2 , and the S REB Ptarget genes acetyl- CoA carboxylas e, 3 - methyl- 3 - hydroxyglutaryl-CoA reductas e, and low- dens ity lipoprotein receptor in liver andadipos e tis s ue and mRNA concentrations of apolipoproteins A- I, A- II,and C- III in the liver were not different between both groups of pigs .In conclus ion, this s tudy s hows that clofibrate treatment activatesPPAR-� in liver and adipos e tis s ue and has a s trong hypotriglyceri-demic and hypocholes terolemic effect in pigs . The finding that mRNAconcentrations of s ome proteins res pons ible for the hypolipidemicaction of fibrates in humans were not altered s ugges ts that there werecertain differences in the mode of action compared with humans . It isals o s hown that PPAR-� activation by clofibrate does not affecthepatic expres s ion of S REB P target genes involved in s ynthes is oftriglycerides and choles terol homeos tas is in liver and adipos e tis s ue ofpigs .

peroxis ome proliferator- activated receptor-� ; choles terol; triglycer-ides

FIB RATES ARE A GROUP OF HYPOLIPIDEMIC agents that have been inclinical us e for s everal decades in humans ( 46) . It is welles tablis hed that thes e agents act as s ynthetic agonis ts of per-oxis ome proliferator- activating receptor-� ( PPAR-�) , a nu-clear receptor als o activated by natural ligands s uch as freefatty acids or s ome eicos anoids . PPAR-� is an importantregulator of cellular fatty acid uptake and intracellular fattyacid trans port, mitochondrial and peroxis omal fatty acid oxi-dation, ketogenes is , and gluconeogenes is ( 48 ) . In humans , themos t pronounced effect of fibrates is a decreas e in plas matriglyceride- rich lipoproteins . Concentrations of low- dens itylipoprotein ( LDL) choles terol generally decreas e in individualswith elevated bas eline plas ma concentrations , and plas mahigh- dens ity lipoprotein ( HDL) choles terol concentrations are

us ually increas ed when bas eline concentrations are low ( 46) .Effects of PPAR-� activation have been mos tly s tudied inrodents , which exhibit a s trong expres s ion of PPAR-� in liverand s how peroxis ome proliferation in the liver in res pons e toPPAR-� activation ( 3 6) . Expres s ion of PPAR-� and s ens itivityto peroxis omal induction by PPAR-� agonis ts , however, varygreatly among s pecies ( 1 9 , 21 ) . In contras t to rats and mice,which are highly s ens itive to induction by peroxis ome prolif-erators , guinea pigs , monkeys , pigs , and humans are relativelyins ens itive ( 21 , 3 2, 3 9 , 48 ) . In thes e nonproliferating s pecies ,expres s ion of PPAR-� in the liver is much lower and theres pons e of many genes to PPAR-� activation is weaker thanin proliferating s pecies ( 8 ) .

In contras t to rodents , there is little information to date aboutthe effects of PPAR-� agonis ts on lipid metabolis m in pigs ,which are not only of agricultural importance but are als o avaluable model for s tudying the lipid metabolis m becaus e oftheir clos e relations hip to humans ( 6) . It has been s hown thattreatment of pigs with clofibrate s timulates mitochondrial andperoxis omal � - oxidation in liver, mus cle, and kidney ( 3 4, 5 6) .Moreover, it was found that pigs expres s functional PPAR-� inthe liver, and s everal target genes induced in the liver byPPAR-� activation have been identified ( 8 ) . However, effectsof PPAR-� activation on lipid concentrations in plas ma andliver of pigs have not yet been inves tigated. In contras t to rats ,mice, or humans in which PPAR-� is predominant in liver( 1 1 ) , pigs exhibit als o a high expres s ion of PPAR-� in adipos etis s ue, and it has been s ugges ted that pigs have a cons iderablecapacity for � - oxidation in adipos e tis s ue ( 1 3 ) . The effect ofPPAR-� agonis ts on gene expres s ion of PPAR-� target genesin adipos e tis s ue of pigs , however, has not yet been inves ti-gated.

Recent s tudies ( 1 7 , 23 , 25 , 3 3 ) in rodents s ugges ted thatactivation of PPAR-� influences hepatic triglyceride s ynthes isand choles terol homeos tas is by interacting with gene expres -s ion or proteolytic activation of s terol regulatory element-binding proteins ( S REB Ps ) , key regulators of lipid s ynthes isand homeos tas is . S REB P- 1 preferentially activates genes re-quired for fatty acid s ynthes is , whereas S REB P- 2 preferen-tially activates the LDL receptor gene and various genesrequired for choles terol s ynthes is ( 22) . S REB Ps are s ynthe-s ized as inactive precurs ors bound to the endoplas matic retic-ulum membranes . For activation to occur, membranes have tobe cleaved by two res ident proteas es within the Golgi, whichs equentially cleave the S REB Ps and releas e the amino- terminalbHLH- Zip- containing domain from the membrane, allowing itto trans locate to the nucleus and activate trans cription of target

Addres s for reprint reques ts and other corres pondence: K. Eder, Ins titutfur Agrar- und Ernahrungs wis s ens chaften, Martin- Luther- Univers itat Halle-Wittenberg, Emil- Abderhalden- S tr. 26, D- 061 08 Halle ( S aale) , Germany( e- mail: Klaus . eder@ landw. uni- halle. de) .

The cos ts of publication of this article were defrayed in part by the paymentof page charges . The article mus t therefore be hereby marked “advertisement”in accordance with 1 8 U. S . C. S ection 1 7 3 4 s olely to indicate this fact.

Am J Physiol Regul Integr Comp Physiol 29 3 : R000 –R000, 2007 .Firs t publis hed March 1 5 , 2007 ; doi: 1 0. 1 1 5 2/ajpregu. 00603 . 2006.

03 63 - 61 1 9 /07 $ 8 . 00 Copyright © 2007 the American Phys iological S ocietyhttp: //www. ajpregu. org R1

AQ: 1

AQ: 2

AQ: 3

AQ: 4

AQ: 12

AQ: 5

AQ: 12

tap rai d4/zh6- areg/zh6- areg/zh600707/zh65828d07a xppws S� 1 4/20/07 9: 48 MS: R- 00603- 2006 I n i : 07/rgh /dh

3. Originalarbeiten

16

genes . Ins ulin- induced genes (Insig) -1 and Insig-2 are modu-lators of S REB P activity ( 5 3 , 5 5 ) . They block the proteolyticcleavage and trans criptional activation of S REB P. In pigs , incontras t to rodents in which lipogenes is takes place primarilyin the liver ( 1 5 , 44) , adipos e tis s ue is the major s ite oflipogenes is . It has been s hown that S REB P- 1 and its targetgene fatty acid s ynthas e, one of the key enzymes of de novofatty acid s ynthes is , are expres s ed at a higher level in pigadipos e tis s ue than in pig liver ( 1 2, 1 3 ) . Whether a link exis tsals o in pigs between PPAR-� activation and gene expres s ion orproteolytic activation of S REB Ps in liver or adipos e tis s ue,which in turn could influence lipid s ynthes is , is pres entlyunknown.

The objective of the pres ent s tudy was to inves tigate theeffects of clofibrate treatment on expres s ion of genes involvedin lipid metabolis m in liver and adipos e tis s ue of pigs . We wereparticularly interes ted in to what extent clofibrate upregulatesPPAR-� target genes in liver and adipos e tis s ue of pigs andwhether there is an interaction between PPAR-� activation andgene expres s ion or proteolytic activation of S REB Ps in thes etis s ues . Therefore, we performed an experiment in which pigswere treated with clofibrate. We us ed relatively young pigswith a body weight s lightly in exces s of 1 0 kg becaus e it hasbeen recently s hown that s uch young pigs expres s a functionalPPAR-� in the liver ( 8 ) . To characterize hepatic PPAR-�expres s ion in the pig model, we compared mRNA concentra-tion of PPAR-� in pig liver with thos e of rat and human livers .To as s es s PPAR-� expres s ion in pig adipos e tis s ue, we com-pared PPAR-� mRNA concentration in pig liver and pigadipos e tis s ue. To examine PPAR-� activation in this animalmodel by clofibrate, we determined mRNA concentrations ofs everal PPAR-� target genes in liver and adipos e tis s ue andals o plas ma concentration of 3 - hydroxybutyrate becaus e it isknown that PPAR-� activation leads to s timulation of keto-genes is ( 28 ) . To find out whether activation of PPAR-� byclofibrate treatment als o affects expres s ion or proteolytic pro-ces s ing of S REB Ps in pig liver or adipos e tis s ue, we deter-mined gene expres s ion of S REB P- 1 , S REB P- 2, Insigs, andtarget genes of S REB P- 1 [ acetyl- CoA carboxylas e ( ACC) ] andS REB P- 2 [ 3 - hydroxy- 3 - methylglutaryl- CoA ( HMG- CoA) re-ductas e and LDL receptor] in thes e tis s ues . To explore themolecular bas is of alterations of plas ma lipoprotein concentra-tions , we als o determined genes involved in lipoprotein me-tabolis m s uch as apolipoproteins ( apo) A- I, apoA- II, and apoC-III and micros omal triglyceride trans fer protein ( MTP) .

MATERIALS AND METHODS

Animals and treatments. Eighteen male 8 - wk- old cros s bred pigs[ ( German Landrace � Large White) � Pietrain] were kept in a roomunder controlled temperature at 23 � 2° C and 5 5 � 5 % relativehumidity with lights on from 0600 to 1 8 00. One day before thebeginning of the experimental feeding period, the pigs were weighedand randomly allocated to two groups , with body weights of 1 2. 0 �0. 4 kg in the control group and 1 1 . 9 � 0. 2 kg in the treatment group( means � S E) . B oth groups of pigs received a nutritionally adequatediet ( 3 1 ) for growing pigs , which contained ( in g/kg) 400 wheat, 23 0s oybean meal, 1 5 0 wheat bran, 1 00 barley, and 9 0 s unflower oil, aswell as a mineral premix that included L- lys ine, DL- methionine, andL- threonine ( 3 0) . This diet contained 1 4. 4 MJ metabolizable energyand 1 8 5 g crude protein/kg. The diet of the treatment group wass upplemented with 5 g clofibrate/kg diet. Diet intake was controlled,and each animal in the experiment was offered an identical amount of

diet per day. The amount of diet adminis tered was � 1 5 % below thatcons umed ad libitum by pigs of a s imilar weight ( as as s es s ed in aprevious s tudy) . Therefore, the diet offered was completely taken inby all pigs in the experiment. During the feeding period, the amountof diet offered each day was increas ed continuous ly from 400 to 1 , 200 g.The pigs had free acces s to water via nipple drinking s ys tems . Theexperimental diets were adminis tered for 28 days . All experimentalprocedures des cribed followed es tablis hed guidelines for the care andus e of laboratory animals and were approved by the local veterinaryoffice.Sample collection. After completion of the feeding period, the

animals were killed under light anes thes ia. Four hours before eutha-nas ia, each pig was fed its res pective diet. After death, blood wascollected into heparinized polyethylene tubes . Plas ma was obtained bycentrifugation of the blood ( 1 , 1 00 g at 4° C for 1 0 min) . Plas malipoproteins were s eparated by s tep- wis e ultracentrifugation ( Mikro-Ultrazentrifuge; S orvall Products , B ad Homburg, Germany) at9 00, 000 g at 4° C for 1 . 5 h. Plas ma dens ities were adjus ted by s odiumchloride and potas s ium bromide, and the lipoprotein fractions � �1 . 006 kg/l [ very low- dens ity lipoproteins ( VLDL) plus chylomicrons ] ,1 . 006 kg/l � � � 1 . 063 kg/l ( LDL) , and � � 1 . 063 kg/l ( HDL) wereremoved by s uction. The liver was dis s ected and weighed, ands amples of liver, s keletal mus cle ( longis s imus dors al mus cle) , ands ubcutaneous adipos e tis s ue ( backfat, at the level of the 1 3 th/1 4th rib)were s tored at � 8 0° C until analys is . For comparis on of PPAR-�expres s ion in rats , humans , and pigs , liver s amples of three male adultrats ( 3 62 � 25 g) and three male adult humans ( collected during ares ection of a tes t s ample for a his topathological evaluation) and liverand adipos e tis s ue of three randomly s elected piglets of the controlgroup were us ed.Lipid analysis. Lipids from liver were extracted with a mixture of

n- hexane and is opropanol ( 3 : 2, vol/vol) ( 1 8 ) . After lipid extracts weredried, aliquots were dis s olved with Triton X- 1 00 ( 1 0) . The concen-trations of choles terol and triglycerides in the lipoprotein fractions ,plas ma, and liver were determined with enzymatic kits ( no.1 1 3 009 9 9 03 1 4 for choles terol and no. 1 5 7 609 9 9 03 1 4 for triglycer-ides , Ecoline S� ; DiaS ys , Holzheim, Germany) .Determination of3-hydroxybutyrate. Concentration of 3 - hydroxy-

butyrate in plas ma was determined with an enzymatic as s ay ( no.1 09 07 9 7 9 03 5 ; R- B IOPHARM, Darms tadt, Germany) .RT-PCR analysis. Total RNAs from liver tis s ue, s keletal mus cle,

and adipos e tis s ue were is olated by a tis s ue lys er ( Qiagen, Hilden,Germany) us ing Trizol reagent ( Invitrogen, Karls ruhe, Germany)according to the manufacturer’ s protocol. RNA concentration andpurity were es timated from optical dens ities at 260 and 28 0 nm( S pectraFluor Plus ; Tecan, Crails heim, Germany) . The quality of allRNA s amples was furthermore as s es s ed by agaros e gel electrophore-s is . Total RNA ( 1 . 2 �g) was us ed for cDNA s ynthes is as des cribedprevious ly ( 24) . The mRNA concentration of genes was meas ured byrealtime PCR, us ing S YB R green I and an MJ Res earch Opticons ys tem ( B iozym Diagnos tik, Oldendorf, Germany) . Real- time PCRwas performed with 1 . 25 U of Taq DNA polymeras e, 5 00 �M dNTPs ,and 26. 7 pmol of the s pecific primers . Amplification efficiencies forall primer pairs were determined by template dilution s eries . Calcu-lation of the relative mRNA concentration was made with the ampli-fication efficiencies and the thres hold cycle values ( 3 7 ) . The hous e-keeping gene GAPDH was us ed for normalization. The PCR primersus ed for real- time RT- PCR were obtained from Operon ( Koln, Ger-many) and Roth ( Karls ruhe, Germany) , res pectively, and are lis ted inTable 1 .Quantification ofPPAR-� mRNA. For the quantification of copy

numbers of PPAR-� mRNA, is olation of total RNA and cDNAs ynthes is from rat, pig, and human liver tis s ues and pig white adipos etis s ue were performed as des cribed above. Real- time PCR was carriedout with s pecific primers ( Table 1 ) as des cribed above; afterward, analiquot of 1 0 � l per PCR product was s ubmitted to agaros e gelelectrophores is to create s tandard templates . After dis s ection from

R2 EFFECTS OF CLOFIB RATE TREATMENT IN PIGS

AJP-Regul Integr Comp Physiol • VOL 29 3 • JULY 2007 • www. ajpregu. org

AQ: 6

AQ: 7

AQ: 8

AQ: 9

AQ: 10

T1 ,AQ: 11


17

ethidium bromide- s tained gel, probes from each s pecies and tis s uewere pooled and eluted with the peqGOLD gel extraction kit ( PeqLabB iotechnologie, Erlangen, Germany) . For the generation of s tandardcurves , the concentration of double- s tranded cDNA was meas ured bya Pico green double- s tranded DNA quantitation kit ( MolecularProbes , Leiden, The Netherlands ) . The calculated molecular weight ofeach PCR product was converted into copy numbers by us ing Avo-gadro’ s number ( 1 mol � 6. 022 � 1 023 molecules ) . The thres holdcycles meas ured by real- time PCR were plotted vs . the copy numbersof diluted s tandard templates to create a s tandard curve. On the bas isof the res pective s tandard curve, the thres hold cycle of each probewas us ed to calculate the copy number of PPAR-� mRNA and wasnormalized to 1 ng of total RNA.Statistics. The res ults were analyzed with Minitab ( S tate College,

PA) s tatis tical s oftware ( releas e 1 3 ) . S tatis tical s ignificances of dif-ferences between control group and treatment group were evaluatedwith S tudent’ s t- tes t. Mean values were cons idered s ignificantlydifferent at P � 0. 05 . Data in the text are pres ented as means � S E.

RESULTS

Comparison ofmRNA concentrations ofPPAR-� in human,rat, and pig liver and in pig adipose tissue. To characterizePPAR-� gene expres s ion in the pig model us ed, we determinedmRNA concentrations of PPAR-� in liver and adipos e tis s ue ofthree control pigs and compared them with thos e in livers ofthree adult rats and three male human s ubjects . PPAR-�mRNA concentrations , corrected for total RNA concentration,were s imilar in human and in pig liver; PPAR-� mRNAconcentrations in human or pig liver were, however, � 1 0- foldlower than concentrations in rat liver ( Fig. 1 ) . In pigs , mRNAconcentrations of PPAR-� were s imilar in adipos e tis s ue and inliver ( Fig. 1 ) .Food intake and body and liver weights in control pigs and

pigs treated with clofibrate. B ecaus e we us ed a controlledfeeding s ys tem, food intake throughout the feeding period was

the s ame for each pig in the experiment, averaging 69 6 � 2g/day. B ody weight after the 28 - day experiment period did notdiffer between control pigs and pigs treated with clofibrate( Table 2) . Relative liver weights , expres s ed per kilogram ofbody weight, were higher in pigs treated with clofibrate than incontrol pigs (P � 0. 05 ; Table 2) .Gene expression in the liver ofcontrol pigs and pigs treated

with clofibrate. mRNA concentration of PPAR-� in liver didnot differ between both groups of pigs ( control: 1 . 00 � 0. 1 3 ;clofibrate: 0. 9 2 � 0. 07 ; n � 9 ) , whereas relative mRNAconcentrations of acyl- CoA oxidas e ( ACO) , carnitine palmi-toyltrans feras e ( CPT) I, liver fatty acid binding protein ( L-FAB P) , mitochondrial HMG- CoA s ynthas e, and s tearoyl- CoAdes aturas e ( S CD) in the liver were moderately increas ed ( 1 . 7 -

Table 1 . Characteristics ofthe specific primers used for RT-PCR

Gene Forward Primer ( from 5 � to 3 � ) Revers e Primer ( from 5 � to 3 � ) bp Annealing Temperature GenB ank No.

ACC C TC C AGGAC AGC AC AGATC A GC C GAAAC ATC TC TGGGATA 1 7 0 60° C AF1 7 5 3 08ACO C TC GC AGAC C C AGATGAAAT TC C AAGC C TC GAAGATGAGT 21 8 60° C AF1 8 5 048apoA- I C GATC AAAGAC AGTGGC AGA GC TGC AC C TTC TTC TTC AC C 23 4 60° C NM_21 43 9 8apoA- II GGAAGGAAGGAAGGAC GAAC TC C C AGAAGTC GGTGAAC TT 1 5 6 60° C AJ5 641 9 6apoC- III GAC AC C TC C C TTC TGGAC AA TC C C AGAAGTC GGTGAAC TT 1 8 5 60° C NM_001 0028 01CPT- I GC ATTTGTC C C ATC TTTC GT GC AC TGGTC C TTC TGGGATA 1 9 8 60° C AF28 8 7 8 9CYP7 TATAGGGC AC GATGC AC AGA AC C TGAC C AGTTC C GAGATG 200 60° C NM_001 005 3 5 2GAPDH AGGGGC TC TC C AGAAC ATC ATC C TC GC GTGC TC TTGC TGGGGTTGG 446 60° C AF01 7 07 9HMG- CoA- R GGTC AGGATGC GGC AC AGAAC G GC C C C AC GGTC C C GATC TC TATG 1 27 65 ° C S 7 9 67 8Ins ig- 1 AGAGGGAGTGGGC C AGTGTGATGC AC GGGAGC C AGGAGC GGATGTAG 27 6 65 ° C AY3 3 6601Ins ig- 2 AAATC AC GC C AGC GC TAAAGTG TC C TAC TC C AAGGC C AAAAC C AC 1 27 60° C AY5 8 5 269LDL- R TGC GAAGATATC GAC GAGTG TAC GGTC C AGGGTC ATC TTC 1 9 6 62° C AF1 1 8 1 47L- FAB P TTC GGTGC ATGTC TAAGC TG TGAGAGGGAGAGGATGAGGA 200 60° C DQ1 8 23 23LPL TGGAC GGTGAC AGGAATGTA AAGGC TGTATC C C AGGAGGT 23 7 60° C NM_21 428 6mHMG- CoA- S GGAC C AAAC AGAC C TGGAGA ATGGTC TC AGTGC C C AC TTC 1 9 8 62° C U9 08 8 4MTP C AGGAC GGC AAAGAAAGAAGG ATGGGAAGC AAAAC C AC AAGG 1 9 9 60° C AY21 7 03 4PPAR-� , rat C C C TC TC TC C AGC TTC C AGC C C C C AC AAGC GTC TTC TC AGC C ATG 5 5 5 65 ° C NM_01 3 1 9 6PPAR� , human TGTGGC TGC TATC ATTTGC TGTGG C TC C C C C GTC TC C TTTGTAGTGC 3 44 60° C NM_001 001 9PPAR� , pig C AGC C TC C AGC C C C TC GTC GC GGTC TC GGC ATC TTC TAGG 3 8 1 60° C DQ43 7 8 8 7S CD AC GTTGTGC C AGTGAGTC AG GTC TTGGC C TC TTGTGC TTC 206 62° C NM_21 3 7 8 1S REB P- 1 C C TC TGTC TC TC C TGC AC C AC AAAGAGAAGC GC C AAGAA 21 3 62° C NM_21 41 5 7S REB P- 2 C GC TC GC GAATC C TGC TGTG GGTGC GGGTC C GTGTC GTG 1 03 65 ° C DQ02047 6

ACO, acyl- CoA oxidas e; ACC, acetyl- CoA carboxylas e; apo, apolipoprotein; CPT- 1 , carnitine palmitoyltrans feras e 1 ; CYP7 , choles terol- 7�- hydroxylas e;HMG- CoA- R, 3 - hydroxy- 3 - methylglutaryl- CoA reductas e; lns ig- 1 and - 2; ins ulin- induced genes 1 and 2; LDL- R, low- dens ity lipoprotein receptor; LPL,lipoprotein lipas e; L- FAB P, liver fatty acid binding protein; mHMG- CoA- S , mitochondrial 3 - hydroxy- 3 - methylglutaryl- CoA s ynthas e; MTP, micros omaltriglyceride trans fer protein; PPAR-� , peroxis ome proliferator- activated receptor � ; S REB P- 1 and - 2, s terol regulatory element- binding protein 1 and 2; S CD,s tearoyl- CoA des aturas e. * For s emiquantitative PCR.

Fig. 1 . mRNA concentrations of peroxis ome proliferator- activated receptor( PPAR) -� in liver tis s ue of rats , humans , and control pigs and in adipos e tis s ueof control pigs . mRNA concentrations were determined by real- time quanti-tative PCR and normalized to total RNA concentration. Data are means � S Eof 3 s amples per s pecies .

R3EFFECTS OF CLOFIB RATE TREATMENT IN PIGS


F1

T2


18

to 2. 5 - fold) in pigs treated with clofibrate compared withcontrol pigs (P � 0. 05 ; Fig. 2) . Relative mRNA concentrationsof MTP and apoA- I, apoA- II, and apoC- III were not differentbetween the two groups of pigs ( relative mRNA concentrationsin control pigs and pigs treated with clofibrate were 1 . 00 �0. 1 0 vs . 0. 9 0 � 0. 1 4 for MPT, 1 . 00 � 0. 1 5 vs . 1 . 1 0 � 0. 1 0 forapoA- I, 1 . 00 � 0. 1 7 vs . 0. 8 6 � 0. 1 1 for apoA- II, and 1 . 00 �0. 1 9 vs . 0. 8 5 � 0. 1 8 for apoC- III, res pectively; n � 9 foreach group) . Relative mRNA concentrations of S REB P- 1 ,S REB P- 2, Insig-1 , and Insig-2 in the liver did als o not differbetween both groups of pigs ( relative mRNA concentrations incontrol pigs and pigs treated with clofibrate were 1 . 00 � 0. 1 0vs . 0. 9 3 � 0. 1 3 for S REB P- 1 , 1 . 00 � 0. 20 vs . 1 . 1 6 � 0. 27 forS REB P- 2, 1 . 00 � 0. 1 6 vs . 0. 9 7 � 0. 1 5 for Insig-1 , and 1 . 00 �0. 20 vs . 1 . 1 8 � 0. 21 for Insig-2 , res pectively; n � 9 for eachgroup) . Hepatic mRNA concentrations of ACC, a target geneof S REB P- 1 , and HMG- CoA reductas e as well as LDL recep-tor, target genes of S REB P- 2, were als o not different betweenboth groups ( relative mRNA concentrations in control pigs and

pigs treated with clofibrate were 1 . 00 � 0. 09 vs . 0. 8 2 � 0. 1 4for ACC, 1 . 00 � 0. 1 2 vs . 0. 8 4 � 0. 1 0 for HMG- CoA reduc-tas e, and 1 . 00 � 0. 1 1 vs . 0. 8 8 � 0. 1 2 for LDL receptor,res pectively; n � 9 for each group) . mRNA concentrations ofcholes terol- 7�- hydroxylas e ( CYP7 ) in liver ( control: 1 . 00 �0. 09 ; clofibrate: 1 . 03 � 0. 1 3 ; n � 9 ) and of lipoprotein lipas ein mus cle ( control: 1 . 00 � 0. 1 3 ; clofibrate: 1 . 1 3 � 0. 1 5 ; n �9 ) were als o not different between both groups of pigs . mRNAconcentrations of lipoprotein lipas e in the liver were notdetectable by mRNA analys is in pigs of both groups .Gene expression in adipose tissue ofcontrol pigs and pigs

treated with clofibrate. mRNA concentration of PPAR-� inadipos e tis s ue did not differ between both groups of pigs( control: 1 . 00 � 0. 1 7 ; clofibrate: 1 . 1 2 � 0. 1 6; n � 9 ) , whereaspigs treated with clofibrate had moderately increas ed mRNAconcentrations of the PPAR-� target genes ACO, CPT- 1 , andS CD (P � 0. 05 ; Fig. 3 ) . mRNA concentration of lipoproteinlipas e, another PPAR-� target gene, in adipos e tis s ue was notdifferent between both groups ( control: 1 . 00 � 0. 1 1 ; clofibrate:0. 9 0 � 0. 1 7 ; n � 9 ) . mRNA concentrations of S REB P- 1 ,S REB P- 2, Insig-1 , and Insig-2 , and S REB P downs tream genesACC, HMG- CoA reductas e, and LDL receptor in adipos etis s ue did not differ between both groups of pigs ( relativemRNA concentrations in control pigs and pigs treated withclofibrate were 1 . 00 � 0. 06 vs . 1 . 1 0 � 0. 08 for S REB P- 1 ,1 . 00 � 0. 1 4 vs . 0. 8 7 � 0. 1 6 for S REB P- 2, 1 . 00 � 0. 1 8 vs .0. 9 4 � 0. 1 3 for Insig-1 , 1 . 00 � 0. 20 vs . 0. 9 2 � 0. 1 6 forInsig-2 , 1 . 00 � 0. 1 9 vs . 1 . 1 9 � 0. 1 8 for ACC, 1 . 00 � 0. 1 0 vs .1 . 1 0 � 0. 1 4 for HMG- CoA reductas e, and 1 . 00 � 0. 08 vs .1 . 07 � 0. 07 for LDL receptor, res pectively; n � 9 for eachgroup) .Concentration of7� -hydroxybutyrate in plasma ofcontrol

pigs and pigs treated with clofibrate. Pigs treated with clofi-brate had a higher concentration of 7� - hydroxybutyrate inplas ma than control pigs ( control: 0. 5 2 � 0. 09 mmol/l; clofi-brate: 2. 1 7 � 0. 1 8 mmol/l; n � 9 , P � 0. 05 ) .Concentrations of triglycerides and cholesterol in liver

plasma and lipoproteins ofcontrol pigs and pigs treated withclofibrate. Pigs treated with clofibrate had lower concentrationsof triglycerides in plas ma and triglyceride rich- lipoproteins

Fig. 2. Relative mRNA concentrations of acyl- CoA oxidas e ( ACO) , carnitinepalmitoyltrans feras e I ( CPT- 1 ) , liver fatty acid binding protein ( L- FAB P) ,mitochondrial 3 - hydroxy- 3 - methylglutaryl- CoA s ynthas e ( mHMG- CoA- S ) ,and s tearoyl- CoA des aturas e ( S CD) in the liver of pigs fed a control diet or adiet s upplemented with 5 g clofibrate/kg diet for 28 days . mRNA concentra-tions were determined by real- time RT- PCR and normalized to GAPDH. Dataare means � S E of 9 animals per group and are expres s ed relative to mRNAconcentrations of control pigs (� 1 ) . * S ignificantly different from controlgroup (P � 0. 05 ) .

Fig. 3 . Relative mRNA concentrations of ACO, CPT- 1 , and S CD in theadipos e tis s ue of pigs fed a control diet or a diet s upplemented with 5 gclofibrate/kg diet for 28 days . mRNA concentrations were determined byrealtime RT- PCR and normalized to GAPDH. Data are means � S E of 9animals per group and are expres s ed relative to mRNA concentrations ofcontrol pigs (� 1 ) . * S ignificantly different from control group (P � 0. 05 ) .

Table 2. Body and liver weights and concentrations oftriglycerides and cholesterol in plasma, lipoprotein fractions,and liver tissue in pigs fed a control diet or a dietsupplemented with 5 g clofibrate/kg diet for 28 days

Control Clofibrate

Initial body weight, kg 1 2. 0� 0. 4 1 1 . 9� 0. 2Final body weight, kg 26. 0� 0. 5 25 . 2� 0. 4Liver weight, g/kg body wt 25 . 9� 0. 8 3 0. 9� 0. 9 *Triglycerides

Plas ma, mmol/l 1 . 09� 0. 06 0. 7 8� 0. 08 *VLDL � chylomicrons , mmol/l 0. 9 3� 0. 06 0. 67� 0. 07 *Liver, �mol/g 9 0. 8� 6. 7 8 2. 7� 8 . 1

Choles terolPlas ma, mmol/l 2. 8 3� 0. 08 0. 9 2� 0. 1 3 *LDL, mmol/l 0. 9 7� 0. 05 0. 3 8� 0. 08 *HDL, mmol/l 1 . 1 3� 0. 04 0. 3 1 � 0. 08 *Liver, �mol/g 69 . 0� 3 . 6 7 3 . 1 � 3 . 0

Values are means � S E with 9 animals per group. VLDL, very low- dens itylipoprotein; HDL, high- dens ity lipoprotein. *P � 0. 05 compared with controlgroup.



F2

F3


19

( VLDL � chylomicrons ) than control pigs (P � 0. 05 ) ; triglyc-eride concentrations in the liver did not differ between bothgroups of pigs ( Table 2) . Pigs treated with clofibrate als o hadlower concentrations of total choles terol in plas ma and lowerLDL and HDL levels than control pigs (P � 0. 05 ) ; choles terolconcentrations in liver, however, did not differ between the twogroups of pigs ( Table 2) .

DISCUSSION

To s tudy the effect of clofibrate treatment on lipid metabo-lis m and gene expres s ion in pigs , we performed an experimentwith young pigs . As in other s tudies dealing with the effects ofclofibrate on metabolis m in experimental animals , we addedclofibrate to the diet. The concentration of clofibrate in the dietof 5 g/kg diet was adopted from other s tudies with pigs ( 8 , 3 4,5 6) , res ulting in a daily dos e of 220 mg/kg body wt. This dos eis relatively high compared with dos es us ed in humans fortreatment of hyperlipidemia, which are us ually in the rangebetween 25 and 3 0 mg/kg body wt.

The pres ent s tudy confirms that expres s ion of PPAR-� in pigliver is much lower than that in rat liver. In the young controlpigs , PPAR-� abundance in the liver was � 1 0- fold lower thanthat in the rat liver. The finding that hepatic PPAR-� abun-dance was s imilar in the liver of the pigs us ed in this s tudy asin liver of adult humans indicates that young pigs may be aus eful model for s tudying the res pons e of PPAR-� activation.The finding that PPAR-� mRNA concentrations in humanlivers are much lower than concentrations in rat liver als oagrees with literature data. In the s tudy of Tugwood et al. ( 5 0) ,the abundance of PPAR-� trans cript in human liver vs . that inrat liver was 1 : 5 , which is clos e to the ratio of 1 : 1 0 found in thepres ent s tudy.

The finding that mRNA concentrations of the PPAR-� targetgenes ACO, CPT- 1 , L- FAB P, mitochondrial HMG- CoA s yn-thas e, and S CD in the liver were increas ed by 5 0 –1 5 0%compared with res ults s hown in control animals clearly indi-cates that clofibrate treatment caus ed PPAR-� activation in theliver of the pigs . This is confirmed by an increas ed concentra-tion of 3 - hydroxybutyric acid, indicative of a s timulation ofhepatic ketogenes is , which is a typical res pons e of PPAR-�activation ( 28 ) . The finding that clofibrate caus es a moderateupregulation of PPAR-� target genes agrees well with recents tudies conducted in piglets that were treated with clofibrate. Inthes e s tudies , mRNA concentrations and activities of ACO andCPT- 1 were two to four times higher in livers of piglets treatedwith clofibrate in dos es s imilar to thos e us ed in the pres ents tudy than in untreated piglets ( 3 4, 5 6) . Interes tingly, in ours tudy, clofibrate treatment caus ed a s ignificant increas e in liverweights of pigs by � 1 5 % , indicative of moderate peroxis omeproliferation. Therefore, the pres ent s tudy s ugges ts that clofi-brate not only upregulated PPAR-� target genes in the liver butals o caus ed a moderate peroxis ome proliferation in the pigs . Its hould be noted that upregulation of PPAR-� target genes wasmuch lower than that s hown in rodents , where treatment withPPAR-� agonis ts typically increas es mRNA concentrations ofACO 1 0- to 20- fold compared with untreated controls ( 1 4, 20,23 , 25 ) . The reas on for the comparatively low upregulation ofthes e enzymes in pigs by clofibrate might be the lower hepaticPPAR-� expres s ion in pigs compared with rodents . Further-more, the pres ence of an alternative s pliced PPAR-� is oform,

which lacks the ligand- binding domain, could contribute to thelower res pons ivenes s of the pig to clofibrate ( 49 ) .

It has been found that pigs , in contras t to humans or rodents ,have a high concentration of PPAR-� in adipos e tis s ue. Dinget al. ( 1 2, 1 3 ) found that PPAR-� mRNA concentration,corrected for 1 8 S ribos omal RNA, was three to four timeshigher in s ubcutaneous adipos e tis s ue than in liver of youngpigs with a body weight of 3 0 kg. In the young pigs us ed in thepres ent s tudy, expres s ion of PPAR-� in adipos e tis s ue, cor-rected for total mRNA content, was at a level in s ubcutaneousadipos e tis s ue s imilar to that s hown in liver. This als o confirmsthat pig adipos e tis s ue has a comparatively high expres s ion ofPPAR-� . To our knowledge, this is the firs t s tudy that inves -tigated the effect of treatment with a PPAR-� agonis t onexpres s ion of PPAR-� target genes in adipos e tis s ue of pigs .Our s tudy s hows that the PPAR-� target genes ACO, CPT- 1 ,and S CD are indeed s ignificantly upregulated by clofibrate,although only to a moderate extent. Although we did notperform a direct activation as s ay, we conclude that PPAR-� inadipos e tis s ue is functional and is activated by PPAR-� ago-nis ts . In this s tudy, we did not directly determine fatty acidoxidation in adipos e tis s ue. The finding that genes involved inmitochondrial and peroxis omal were upregulated by clofibrates ugges ts that PPAR-� agonis ts indeed could s timulate � - oxi-dation in pig adipos e tis s ue, which s hould be inves tigated infuture s tudies .

To elucidate a pos s ible link between PPAR-� activation andS REB P- mediated lipid homeos tas is in pigs , we determinedrelative mRNA concentrations of S REB P-1 and -2, Insig-1 and -2,and s ome target genes of S REB P- 1 ( ACC) and S REB P- 2( HMG- CoA reductas e, LDL receptor) in the liver and in theadipos e tis s ue, which is the major s ite of lipogenes is in the pig.We found that mRNA concentrations of S REB P- 1 and - 2,Insig-1 and -2 , and target genes of S REB P- 1 and - 2 in liver andadipos e tis s ue were not different in clofibratetreated andcontrol pigs . This indicates that activation of PPAR-� byclofibrate did not influence expres s ion and activity of S REB P- 1and S REB P- 2 in both tis s ues . S CD is another target gene ofS REB P- 1 involved in lipogenes is , which als o has a PPARres pons e element in its promoter and is upregulated byPPAR-� activation ( 3 0) . We as s ume that an upregulation ofS CD in liver and adipos e tis s ue of pigs treated with clofibratewas probably due to PPAR-� activation. An upregulation ofS CD in the liver of pigs by treatment with clofibrate has als obeen obs erved by Cheon et al. ( 8 ) . S REB P- 1 controls fatty acidand triglyceride s ynthes is , and S REB P- 2 controls choles terols ynthes is and choles terol uptake in cells via the LDL receptor( 22) . It is as s umed that thes e S REB P- controlled proces s es werenot altered by PPAR-� activation in liver and adipos e tis s ue.This as s umption is in contras t to recent findings in mice inwhich WY- 1 4, 643 , a s ynthetic PPAR-� agonis t, s timulatedS REB P- 1 - mediated fatty acid s ynthes is in the liver ( 23 ) andfindings in rats and hams ters in which fibrates decreas edS REB P-2-mediated expres s ion of HMG-CoA reductas e and LDLreceptor ( 1 7 , 25 ) .

The pres ent s tudy s hows for the firs t time that clofibratetreatment s trongly reduces triglyceride concentrations inplas ma and triglyceride- rich lipoproteins in pigs . Thes e res ultsagree with obs ervations in humans and rodents ( 2, 1 6) . S tudiesin humans and rodents have s hown that this effect is in part dueto increas ed oxidation of fatty acids in the liver, leading to




20

reduced triglyceride s ynthes is and s ecretion, and in part due toincreas ed lipolys is caus ed by induction of lipoprotein lipas eactivity and repres s ed trans cription of apoC- III in the liver ( 1 4,1 7 , 41 , 5 4) . The obs ervation that the PPAR-� target genesL- FAB P, ACO, and CPT- 1 were increas ed indicates that clo-fibrate increas ed the uptake of fatty acids into hepatocytes ands timulated both peroxis omal and mitochondrial � - oxidation inpigs . To find out whether clofibrate s timulates lipolys is , wedetermined mRNA concentrations of lipoprotein lipas e in liver,adipos e tis s ue, and mus cle, extrahepatic tis s ues that als o ex-pres s comparatively high levels of PPAR-� ( 4) , and mRNAconcentration of apoC- III, an inhibitor of lipoprotein lipas e.The obs ervation that lipoprotein lipas e mRNA could not bedetected in liver of pigs of both groups s ugges ts that the pig hasgenerally a low expres s ion of lipoprotein lipas e in the liver.This s ugges tion is confirmed by recent s tudies that als o s howeda very low gene expres s ion of lipoprotein lipas e in liver of pigs( 27 , 44) . The finding that clofibrate does not reduce apoC- IIIagrees with obs ervations in monkeys ( 42) but dis agrees withobs ervations in human and rat hepatocytes and livers of ham-s ters in which apoC- III expres s ion was downregulated byfibrates ( 9 , 1 7 , 26) . B ecaus e lipoprotein lipas e in adipos e tis s ueand mus cle was als o not upregulated, the data of this s tudy donot give any indication of increas ed lipolys is in pigs treatedwith clofibrate. It has recently been s hown in monkeys thatadminis tration of a PPAR-� agonis t increas es s erum levels ofapoA- V, which has recently been recognized as a key regulatorof s erum triglyceride concentrations ( 42) . Overexpres s ion ofapoA- V in mice caus ed a s trong reduction of s erum triglycer-ide concentrations ( 3 5 , 5 1 ) , and upregulation of apoA- V couldbe involved in the hypotriglyceridemic effect of PPAR-�agonis ts . For technical reas ons , we were unable to determinegene expres s ion of apoA- V, but it is pos s ible that clofibratereduced triglyceride concentrations in pigs by upregulation ofapoA- V.

In this s tudy, we als o determined hepatic mRNA concentra-tions of MTP, the rate- limiting protein for as s embly ands ecretion of VLDL in the liver, which has recently beendemons trated in mice to be a PPAR-� target gene ( 1 ) . Ours tudy s hows that activation of PPAR-� by clofibrate does nots timulate gene expres s ion of MTP in pigs , in contras t to rats ,and probably does not s timulate s ecretion of lipids from theliver into the blood via VLDL. This could help to explain theobs ervation that concentrations of triglycerides and choles terolin the liver remained unchanged in pigs after clofibrate treat-ment.

In humans , treatment with fibrates us ually reduces plas maand LDL choles terol concentrations and increas es HDL cho-les terol ( 46) . The pres ent s tudy s hows that clofibrate reducesplas ma and LDL choles terol concentrations in pigs , as als os hown in humans . The effect of clofibrate on HDL choles terol,however, is oppos ite to that obs erved in humans . The elevationof HDL choles terol by fibrate treatment in humans is caus edprimarily by increas ed gene expres s ion of apoA- I and apoA- IIin the liver ( 3 , 5 2) . In contras t to humans , fibrates lower plas maHDL concentrations in rats and hams ters becaus e of a decreas eof liver apoA- I and apoA- II gene expres s ion ( 1 7 , 45 , 47 ) . Thefinding that mRNA concentrations of apoA- I and apoA- II inthe liver did not differ between both groups of pigs s ugges tsthat clofibrate reduced HDL choles terol by a mechanis m otherthan HDL reduction.

It has been s hown in mice that activation of PPAR-� leadsto downregulation of CYP7 , the key enzyme of hepatic bileacid formation, which is probably becaus e of reduced avail-ability of hepatic nuclear factor- 4, a trans cription factor in-volved in the bas al expres s ion of CYP7 ( 29 , 3 8 ) . The pres ents tudy s hows that activation of PPAR-� by clofibrate does notalter hepatic mRNA concentrations of CYP7 , which, alongwith LDL receptor and HMG- CoA reductas e, is a key factor ofhepatic choles terol homeos tas is ( 40) . The finding that mRNAconcentrations of thes e three genes were not altered by clofi-brate treatment agrees with a recent s tudy in which pigs weretreated with clofibrate ( 3 4) and is in accordance with theobs ervation that hepatic choles terol concentrations were als ounchanged in pigs treated with clofibrate compared with con-trol pigs . Thes e data als o s ugges t that the greatly reducedconcentration of LDL choles terol is not becaus e of a loweredcholes terol s ynthes is , enhanced elimination of choles terol fromthe liver via bile acid formation, or upregulation of LDLreceptor. It has been s hown in humans that fibrate treatmentenhances LDL uptake via the LDL receptor, not as a res ult ofincreas ed LDL receptor expres s ion but as a res ult of theformation of LDL particles with a higher affinity to the LDLreceptor ( 5 , 7 ) . It is pos s ible that a s imilar effect is res pons iblefor the s trongly reduced LDL levels in pigs treated withclofibrate. This s hould be inves tigated further in future s tudies .

In conclus ion, this s tudy s hows that clofibrate treatmentcaus es an increas e in liver weights , indicative of peroxis omeproliferation, and moderate upregulation of PPAR-� targetgenes involved in liver and adipos e tis s ue. Clofibrate treatmentcaus es a s trong reduction of triglyceride and choles terol con-centrations in plas ma and lipoproteins , which agrees withfindings in rodents . Gene expres s ion analys is of lipoproteinlipas e, apoA- I, apoA- II, and apoC- III in the liver s ugges ts thatbiochemical mechanis ms underlying thes e effects might be inpart different from thos e in humans or rodents . In pigs , unlikerodents , hepatic concentrations of triglycerides and choles terolare not altered by clofibrate. We als o s howed that PPAR-�activation by clofibrate does not affect expres s ion of S REB Ptarget genes , which are involved in s ynthes is of triglyceridesand choles terol, in liver and adipos e tis s ue of pigs .

ACKNOWLEDGMENTS

The authors thank B ettina Konig for critical dis cus s ion of the manus cript.

REFERENCES

1 . Ameen C, Edvardsson U, Ljungberg A, Asp L, Akerblad P, Tuneld A,Olofsson SO, Linden D, Oscarsson J. Activation of peroxis ome prolif-erator- activated receptor alpha increas es the expres s ion and activity ofmicros omal triglyceride trans fer protein in the liver. J Biol Chem 28 0:1 224 –1 229 , 2005 .

2. Auwerx J, Schoonjans K, Fruchart JC, Staels B. Regulation of triglyc-eride metabolis m by PPARs : fibrates and thiazolidinediones have dis tincteffects . J Atheroscler Thromb 3 : 8 1 – 8 9 , 1 9 9 6.

3 . Berthou L, Duverger N, Emmanuel F, Langouet S, Auwerx J, Guil-louzo A, Fruchart JC, Rubin E, Denefle P, Staels B, Branellec D.

Oppos ite regulation of human vers us mous e apolipoprotein A- I by fibratesin human apolipoprotein A- I trans genic mice. J Clin Invest 9 7 : 2408 –241 6, 1 9 9 6.

4. Braissant O, Foufelle F, Scotto C, Dauca M, Wahli W. Differentialexpres s ion of peroxis ome proliferator- activated receptors ( PPARs ) : tis s uedis tribution of PPAR- alpha, - beta, and - gamma in the adult rat. Endocri-nology 1 3 7 : 3 5 4 –3 66, 1 9 9 6.

5 . Bruckert E, Dejager S, Chapman MJ. Ciprofibrate therapy normalis esthe atherogenic low- dens ity lipoprotein s ubs pecies profile in combinedhyperlipidemia. Atherosclerosis 1 00: 9 1 –1 02, 1 9 9 3 .




21

6. Carey GB. The s wine as a model for s tudying exercis e- induced changesin lipid metabolis m. Med Sci Sports Exerc 29 : 1 43 7 –1 443 , 1 9 9 7 .

7 . Caslake MJ, Packard CJ, Gaw A, Murray E, Griffin BA, VallanceBD, Sheperd J. Fenofibrate and LDL metabolic heterogeneity in hyper-choles terolemia. Arterioscler Thromb 1 3 : 7 02–7 1 1 , 1 9 9 3 .

8 . Cheon Y, Nara TY, Band MR, Beever JE, Wallig MA, Nakamura MT.Induction of overlapping genes by fas ting and a peroxis ome proliferator inpigs : evidence of functional PPAR alpha in nonproliferating s pecies . Am JPhysiol Regul Integr Comp Physiol 28 8 : R1 5 25 –R1 5 3 5 , 2005 .

9 . Clavey V, Copin C, Mariotte MC, Bauge E, Chinetti G, Fruchart J,Fruchart JC, Dallongeville J, Staels B. Cell culture conditions determineapolipoprotein CIII s ecretion and regulation by fibrates in human hepa-toma HepG2 cells . Cell Physiol Biochem 9 : 1 3 9 –1 49 , 1 9 9 9 .

1 0. De Hoff JL, Davidson JH, Kritchevsky V. An enzymatic as s ay fordetermining free and total choles terol in tis s ues . Clin Chem 24: 43 3 – 43 5 ,1 9 7 8 .

1 1 . Desvergne B, Wahli W. Peroxis ome proliferator- activated receptors :nuclear control of metabolis m. Endocr Rev 20: 649 – 68 8 , 1 9 9 9 .

1 2. Ding ST, McNeel RL, Mersmann HJ. Expres s ion of porcine adipocytetrans cripts : tis s ue dis tribution and differentiation in vitro and in vivo.Comp Biochem Physiol B Biochem Mol Biol 1 23 : 3 07 –3 1 8 , 1 9 9 9 .

1 3 . Ding ST, Schinckel AP, Weber TE, Mersman HJ. Expres s ion ofporcine trans cription factors and genes related to fatty acid metabolis m indifferent tis s ues and genetic populations . J Anim Sci 7 8 : 21 27 –21 3 4, 2000.

1 4. Frederiksen KS, Wulff EM, Sauerberg P, Mogensen JP, Jeppesen L,Fleckner J. Prediction of PPAR-� ligand- mediated phys iological changesus ing gene expres s ion profiles . J Lipid Res 45 : 5 9 2– 601 , 2004.

1 5 . Gandemier G, Pascal G, Durand G. In vivo changes in the rates of totallipid and fatty acid s ynthes is in liver and white adipos e tis s ues of male ratsduring pos tweaning growth. Int J Biochem 1 4: 7 9 7 – 8 04, 1 9 8 2.

1 6. Gervois P, Toora IP, Fruchart JC, Staels B. Regulation of lipid andlipoprotein metabolis m by PPAR activators . Clin Chem Lab Med 3 8 :3 –1 1 , 2000.

1 7 . Guo Q, Wang PR, Milot DP, Ippolito MC, Hernandez M, Burton CA,Wright SD, Chao Y. Regulation of lipid metabolis m and gene expres s ionby fenofibrate in hams ters . Biochim Biophys Acta 1 5 3 3 : 220 –23 2, 2001 .

1 8 . Hara A, Radin NS. Lipid extraction of tis s ues with a low toxicity s olvent.Anal Biochem 9 0: 420 – 426, 1 9 7 8 .

1 9 . Hawkins JM, Jones WE, Bonner FW, Gibson GG. The effect ofperoxis ome proliferators on micros omal, peroxis omal, and mitochondrialenzyme activities in the liver and kidney. Drug Metab Rev 1 8 : 441 –5 1 5 ,1 9 8 7 .

20. He WS, Nara TY, Nakamura MT. Delayed induction of delta- 6 anddelta- 5 des aturas es by a peroxis ome proliferators . Biochem Biophys Res

Commun 29 9 : 8 3 2– 8 3 8 , 2002.21 . Holden PR, Tugwood JD. Peroxis ome proliferator- activated receptor � :

role in rodent liver cancer and s pecies differences . J Mol Endocrinol 22:1 – 8 , 1 9 9 9 .

22. Horton JD, Goldstein JL, Brown MS. S REB Ps : activators of thecomplete program of choles terol and fatty acid s ynthes is in the liver.J Clin Invest 1 09 : 1 1 25 –1 1 3 1 , 2002.

23 . Knight BL, Hebbach A, Hauton D, Brown AM, Wiggins D, Patel DD.A role for PPAR� in the control of S REB P activity and lipid s ynthes is inthe liver. Biochem J 3 8 9 : 41 3 – 421 , 2005 .

24. Konig B, Eder K. Differential action of 1 3 - HPODE on PPAR� down-s tream genes in rat Fao and human HepG2 hepatoma cell lines . J NutrBiochem 1 7 : 41 0 – 41 8 , 2006.

25 . Konig B, Koch A, Spielmann J, Hilgenfeld C, Stangl GI, Eder K.Activation of PPAR� lowers s ynthes is and concentration of choles terol byreduction of nuclear S REB P- 2. Biochem Pharmacol 7 3 : 5 7 4 –5 8 5 , 2007 .

26. Lawrence JW, Li Y, Chen S, DeLuca JG, Berger JP, UmbenhauerDR, Moller DE, Zhou G. Differential gene regulation in human vers usrodent hepatocytes by peroxis ome proliferator- activated receptor ( PPAR)� . J Biol Chem 27 6: 3 1 5 21 –3 1 5 27 , 2001 .

27 . Lewis GF, Rader DJ. New ins ights into the regulation of HDL metabo-lis m and revers e choles terol trans port. Circ Res 9 6: 1 221 –1 23 2, 2005 .

28 . Mandard S, Muller M, Kersten S. Peroxis ome proliferator- activatedreceptor � target genes . Cell Mol Life Sci 61 : 3 9 3 – 41 6, 2004.

29 . Marrapodi M, Chiang JY. Peroxis ome proliferator- activated receptoralpha ( PPAR�) and agonis t inhibit choles terol 7�- hydroxylas e gene( CYP7 A1 ) trans cription. J Lipid Res 41 : 5 1 4 –5 20, 2000.

3 0. Miller CW, Ntambi JM. Peroxis ome proliferators induce mous e livers tearoyl- CoA des aturas e 1 gene expres s ion. Proc Natl Acad Sci USA 9 3 :9 443 –9 448 , 1 9 9 6.

3 1 . National Research Council. Nutrient Requirement ofSwine ( 1 0th rev.

ed. ) . Was hington, DC: National Academy of S ciences , 1 9 9 8 .

3 2. Orton TC, Adam HK, Bentley M, Holloway B, Tucker MJ. Clobuzarit:

s pecies differences in the morphological and biochemical res pons e of the

liver following chronic adminis tration. Toxicol Appl Pharmacol 7 3 : 1 3 8 –

1 5 1 , 1 9 8 4.

3 3 . Patel DD, Knight BL, Wiggins D, Humphries SM, Gibbons GF.

Dis turbances in the normal regulation of S REB P- s ens itive genes in PPAR

�- deficient mice. J Lipid Res 42: 3 28 –3 3 7 , 2001 .

3 4. Peffer PL, Lin X, Odle J. Hepatic beta- oxidation and carnitine palmi-

toyltrans feras e I in neonatal pigs after dietary treatments of clofibric acid,

is oproterenol, and medium- chain triglycerides . Am J Physiol Regul Integr

Comp Physiol 28 8 : R1 5 1 8 –R1 5 24, 2005 .

3 5 . Pennacchio LA, Olivier M, Hubacek JA, Cohen JC, Cox DR, Fru-

chart JC, Krauss RM, Rubin EM. An apolipoprotein influencing trig-

lycerides in humans and mice revealed by comparative s equencing.

Science 29 4: 1 69 –1 7 3 , 2001 .

3 6. Peters JM, Cheung C, Gonzalez FJ. Peroxis ome proliferator- activated

receptor-� and liver cancer: where do we s tand? J Mol Med 8 3 : 7 7 4 –7 8 5 ,

2005 .

3 7 . Pfaffl MW. A new mathematical model for relative quantification in

real- time RT- PCR. Nucleic Acids Res 29 : e45 , 2001 .

3 8 . Post SM, Duez H, Gervois PP, Staels B, Kuipers F, Princen HM.

Fibrates s uppres s bile acid s ynthes is via peroxis ome proliferator- activated

receptor-�- mediated downregulation of choles terol 7�- hydroxylas e and

s terol 27 - hydroxylas e expres s ion. Arterioscler Thromb Vasc Biol 21 :

1 8 40 –1 8 45 , 2001 .

3 9 . Reinhart GA, Mahan DC, Lepine AJ, Simmen FA, Moore BE. Dietary

clofibric acid increas es intes tinal fatty acid binding protein activity and

apparent lipid diges tibility in weanling s wine. J Anim Sci 7 1 : 269 3 –269 9 ,

1 9 9 3 .

40. Russell DW. Choles terol bios ynthes is and metabolis m. Cardiovasc Drugs

Ther 6: 1 03 –1 1 0, 1 9 9 2.

41 . Schoonjans K, Peinado-Onsurbe J, Lefebvre AM, Heyman RA, Briggs

M, Deeb S, Staels B, Auwerx J. PPAR� and PPAR� activators direct a

dis tinct tis s ue- s pecific trans criptional res pons e via a PPRE in the lipopro-

tein lipas e gene. EMBO J 1 5 : 5 3 3 6 –5 3 48 , 1 9 9 6.

42. Schultze AE, Alborn WE, Newton RK, Konrad RJ. Adminis tration of

a PPAR� agonis t increas es s erum apolipoprotein A- V levels and the

apolipoprotein A- V/apolipoprotein C- III ratio. J Lipid Res 46: 1 5 9 1 –1 5 9 5 ,

2005 .

43 . Semenkovich CF, Chen SH, Wims M, Luo CC, Li WH, Chan L.

Lipoprotein lipas e and hepatic lipas e mRNA tis s ue s pecific expres s ion,developmental regulation, and evolution. J Lipid Res 3 0: 423 – 43 1 , 1 9 8 9 .

44. Semenkovich CF, Coleman T, Fiedorek FT Jr. Human fatty acids ynthas e mRNA: tis s ue dis tribution, genetic mapping, and kinetics ofdecay after glucos e deprivation. J Lipid Res 3 6: 1 5 07 –1 5 21 , 1 9 9 5 .

45 . Staels B, Auwerx J. Regulation of apo A- I gene expres s ion by fibrates .Atherosclerosis Suppl 1 3 7 : S 1 9 –S 23 , 1 9 9 8 .

46. Staels B, Dallongeville J, Auwerx J, Schoonjans K, Leitersdorf E,Fruchart JC. Mechanis m of action of fibrates on lipid and lipoproteinmetabolis m. Circulation 9 8 : 208 8 –209 3 , 1 9 9 8 .

47 . Staels B, Van Tol A, Andreu T, Auwerx J. Fibrates influence theexpres s ion of genes involved in lipoprotein metabolis m in a tis s ue-s elective manner in the rat. Arterioscler Thromb 1 2: 1 7 9 –1 8 7 , 1 9 9 2.

48 . Stott WT, Yano BL, Williams DM, Barnard SD, Hannah MA, Cies-zlak FS, Herman JR. S pecies - dependent induction of peroxis ome pro-liferation by haloxyfop, an aryloxyphenoxy herbicide. Fundam Appl

Toxicol 28 : 7 1 –7 9 , 1 9 9 5 .49 . Sundvold H, Grindflek E, Lien S. Tis s ue dis tribution of porcine perox-

is ome proliferator- activated receptor � : detection of an alternativelys pliced mRNA. Gene 27 3 : 1 05 –1 1 3 , 2001 .

5 0. Tugwood JD, Holden PR, James NH, Prince RA, Roberts RA. Aperoxis ome proliferator- activated receptor- alpha ( PPAR�) cDNA clonedfrom guinea- pig liver encodes a protein with s imilar properties to themous e PPAR� : implications for s pecies differences in res pons es to per-oxis ome proliferators . Arch Toxicol 7 2: 1 69 –1 7 7 , 1 9 9 8 .

5 1 . Van der Vliet HN, Schaap FG, Levels JH, Ottenhoff R, Looije N,Wesseling JG, Groen AK, Chamuleau RA. Adenoviral overexpres s ionof apolipoprotein A- V reduces s erum levels of triglycerides and choles -terol in mice. Biochem Biophys Res Commun 29 5 : 1 1 5 6 –1 1 5 9 , 2002.




22

5 2. Vu-Dac N, Schoonjans K, Kosykh V, Dallongeville J, Fruchart JC,Staels B, Auwerx J. Fibrates increas e human apolipoprotein A- II expres -s ion through activation of the peroxis ome proliferator- activated receptor.J Clin Invest 9 6: 7 41 –7 5 0, 1 9 9 5 .

5 3 . Yabe D, Brown MS, Goldstein JL. Ins ig-2, a s econd endoplas mic reticulumprotein that binds S CAP and blocks export of s terol regulatory element-binding proteins . Proc Natl Acad Sci USA 99: 1 275 3 –1 275 8 , 2002.

5 4. Yamazaki K, Kuromitsu J, Tanaka I. Microarray analys is of geneexpres s ion changes in mous e liver induced by peroxis ome proliferator-

activated receptor alpha agonis ts . Biochem Biophys Res Commun 29 0:1 1 1 4 –1 1 22, 2002.

5 5 . Yang T, Espenshade PJ, Wright ME, Yabe D, Gong Y, Aebersold R,Goldstein JL, Brown MS. Crucial s tep in choles terol homeos tas is : s terolspromote binding of S CAP to INS IG- 1 , a membrane protein that facilitatesretention of S REB Ps in ER. Cell 1 1 0: 48 9 –5 00, 2002.

5 6. Yu XX, Odle J, Drackley JK. Differential induction of peroxis omalbeta- oxidation enzymes by clofibric acid and as pirin in piglet tis s ues . Am JPhysiol Regul Integr Comp Physiol 28 1 : R1 5 5 3 –R1 5 61 , 2001 .




23

Clofibrate Increases Hepatic Triiodothyronine (T3)- and Thyroxine(T4)-Glucuronosyltransferase Activities and Lowers Plasma T3 and

T4 Concentrations in Pigs

Sebastian Luci, Holger Kluge, Frank Hirche, and Klaus Eder

Institut fur Ernahrungswissenschaften, Martin-Luther-Universitat, Halle-Wittenberg, Germany

Received June 7, 2006; accepted August 4, 2006

ABSTRACT:

In rats, clofibrate acts as a microsomal enzyme inducer and dis-rupts the metabolism of thyroid hormones by increasing hepaticglucuronidation of thyroxine. Whether similar effects occur in thepig has not yet been investigated. This study was performed toinvestigate the effect of clofibrate treatment on metabolism ofthyroid hormones in pigs. To this end, an experiment with 18 pigs,which were assigned to two groups, was performed. One groupreceived a control diet, and the other group was fed the same dietsupplemented with 5 g of clofibrate/kg for 28 days. Pigs treatedwith clofibrate had higher hepatic activities of T3- and T4-UDPglucuronosyltransferases (UGT) and lower concentrations of totaland free T4 and total T3 in plasma than control pigs (P < 0.05). Weightsand histology of the thyroid gland (epithelial height, follicle lumen

diameter) did not differ between the two groups, but pigs treated withclofibrate had higher mRNA concentrations of various genes in thethyroid responsive to thyroid-stimulating hormone (TSH) such as TSHreceptor, sodium iodine symporter, thyroid peroxidase, and cathep-sin B than control pigs (P < 0.05). Pigs treated with clofibrate also hadlower hepatic mRNA concentrations of proteins involved in plasmathyroid hormone transport [thyroxine-binding globulin (P < 0.10),transthyretin (P < 0.05), and albumin (P < 0.05)] and thyroid hormonereceptor �1 (P < 0.05) than control pigs. In conclusion, this studyshows that clofibrate treatment induces a strong activation of T3- andT4-UGT in pigs, leading to increased glucuronidation and markedlyreduced plasma concentrations of these hormones, accompanied bya moderate stimulation of thyroid function.

Fibrates are synthetic agonists of peroxisome proliferator-activatedreceptor-� (PPAR�), a nuclear receptor also activated by naturalligands like free fatty acids or some eicosanoids. Activation ofPPAR� leads to up-regulation of transcription of several genes in-volved mainly in mitochondrial and peroxisomal �-oxidation, keto-genesis, and gluconeogenesis (Mandard et al., 2004). Fibrates havebeen in clinical use as hypolipidemic agents for several decades.Several studies in rodents and cell culture systems have shown thatfibrates, like many other drugs (e.g., phenobarbital, 3-methylchol-antrene, polychlorinated biphenyl, tetrachlorobiphenyl, pregnenolone-16�-carbonitrile, or dexamethasone), induce UDP glucuronosyltrans-ferases (UGT) (Beetstra et al., 1991; Saito et al., 1991; Barter andKlaassen, 1992a,b, 1994; Visser et al., 1993a,b; Jemnitz et al., 2000;Viollon-Abadie et al., 2000; Vansell and Klaassen, 2002). UGT,consisting of UGT1 and UGT2 isoforms, are localized in the endo-plasmatic reticulum of hepatocytes and extrahepatic tissue and belongto the enzymes of phase II metabolism. With broad and overlappingsubstrate specificities, the UGT isoenzymes catalyze the glucuronida-tion of differential functional groups, using UDP-glucuronic acid asthe cofactor (Miners and Mackenzie, 1991; Mackenzie et al., 1997).Thyroid hormones thyroxine (T4) and triiodothyronine (T3) are sub-

strates of hepatic UGT, and glucuronidation of these hormones is themain metabolic pathway for deactivating them (Jemnitz et al., 2000).In rats, several of the drugs acting as inducers of microsomal enzymeshave been shown to produce hypertrophy and hyperplasia of thyroidfollicular cells, most probably through increased deactivation of thy-roid hormones by UGT, leading to a reduction of serum T4 andpossibly T3 (Beetstra et al., 1991; Saito et al., 1991; Barter andKlaassen 1992a, 1994). In mice, in contrast to rats, clofibrate treat-ment did not alter T3- and T4-UGT activities and plasma concentra-tions of thyroid hormones (Viollon-Abadie et al., 1999). These studiesshow species-specific differences in the effects of clofibrate on he-patic thyroid hormone metabolism (i.e., glucuronidation of thyroidhormones).

In rodents, PPAR� agonists not only induce many genes involvedin various metabolic pathways such as �-oxidation, ketogenesis, andgluconeogenesis but also cause severe peroxisome proliferation in theliver, hepatomegaly, and hepatocarcinogenesis (Peters et al., 2005). Incontrast to rodents, PPAR� agonists do not induce peroxisome pro-liferation or tumor in the liver of many other species, such as guineapigs, swine, monkeys, and humans, although they retain a hypotri-glyceridemic effect in these species (Holden and Tugwood, 1999). Innonproliferating species, expression of PPAR� in the liver is muchlower, and the response of many genes to PPAR� activation is weakerthan in proliferating species (Cheon et al., 2005). It is known that

Article, publication date, and citation information can be found athttp://dmd.aspetjournals.org.

doi:10.1124/dmd.106.011379.

ABBREVIATIONS: PPAR�, peroxisome proliferator-activated receptor �; UGT, UDP glucuronosyltransferase(s); T4, thyroxine; T3, 3,3�,5-triiodo-thyronine; TSH, thyroid-stimulating hormone; pNP, p-nitrophenol; UDPGA, UDP-glucuronic acid; RT-PCR, reverse transcriptase polymerase chainreaction; GAPDH, glycerinaldehyde-3-phosphate dehydrogenase; ACO, acyl CoA oxidase; CPT-1, carnitine palmitoyl transferase 1.

0090-9556/06/3411-1887–1892$20.00DRUG METABOLISM AND DISPOSITION Vol. 34, No. 11Copyright © 2006 by The American Society for Pharmacology and Experimental Therapeutics 11379/3146383DMD 34:1887–1892, 2006 Printed in U.S.A.

1887 24

PPAR� activation can modulate metabolizing enzymes of phase I andII biotransformation (Rushmore and Kong, 2002; Zhou et al., 2005).Moreover, it has been shown that some UGT isoforms (UGT1A9,UGT2B4) are PPAR� target genes (Barbier et al., 2003a,b). There-fore, nonproliferating species could respond differently from prolif-erating species to clofibrate with respect to induction of UGT (i.e.,UGT involved in glucuronidation of thyroid hormones). To ourknowledge, the effect of clofibrate on the hepatic thyroid hormonemetabolism has not yet been investigated in vivo in a nonproliferatingspecies.

The aim of our study was to investigate the effects of clofibratetreatment on hepatic thyroid hormone metabolism (i.e., activities ofT3- and T4-UGT in pigs, representing a nonproliferating species).Therefore, as well as determining hepatic activities of T3- and T4-UGT, we also measured plasma concentrations of thyroid hormones,thyroid weights, thyroidal epithelial cell height, and follicle lumendiameter and gene expression levels of several thyroidal genes in-volved in thyroid hormone biosynthesis [thyroid-stimulating hormone(TSH) receptor, sodium iodide symporter, thyroid peroxidase, dualoxidase 2, thyroglobulin, cathepsin B, and type II iodothyroninedeiodinase]. We also investigated the effect of clofibrate on mRNAexpression of genes involved in thyroid hormone transport (transthy-retin, thyroxine-binding globulin, and albumin), peripheral conversionof thyroid hormones (type I iodothyronine deiodinase), and thyroidhormone signaling (thyroid hormone receptor �1) in the liver.

Materials and Methods

Chemicals. Bilirubin, Brij 56, clofibrate, dithiothreitol, p-nitrophenol(pNP), 6-propyl-2-thiouracil, cholic acid, T3, T4, Triton X-100, and UDP-glucuronic acid (UDPGA) were obtained from Sigma (Deisenhofen, Ger-many); 125I-T3 (3076 �Ci/�g) and 125I-T4 (1500 �Ci/�g) were obtained fromAmersham Biosciences (Freiburg, Germany); and bicinchoninic acid proteinassay reagent was from Interchim (Montelucon, France).

Animals and Treatments. Eighteen male 8-week-old crossbred [(GermanLandrace � Large White) � Pietrain] pigs, bred in the local animal facility,were used. They weighed between 11.0 and 13.5 kg. They were individuallyhoused in a room maintained at 23°C and 50 to 60% relative humidity withlight from 6:00 AM to 6:00 PM. On the day before the start of the experimentalfeeding period, all the pigs were weighed and assigned to two groups withbody weights of 12.0 � 1.1 (S.D.) kg (control group) and 11.9 � 0.6 (S.D.) kg(treatment group). Both groups of pigs received a nutritionally adequate diet(National Research Council, 1998) for growing pigs containing wheat (400g/kg), soybean meal (230 g/kg), wheat bran (150 g/kg), barley (100 g/kg),sunflower oil (90 g/kg), and mineral premix including L-lysine, DL-methionine,and L-threonine (30). This diet contained 14.4 MJ metabolizable energy and

185 g of crude protein/kg. The diet of the treatment group was supplementedwith 5 g of clofibrate/kg. To standardize feed intake, each pig within theexperiment received 700 g of the diet daily, which was completely consumedby all the animals in the experiment. The clofibrate dosage in the treated pigswas 220 mg/kg b.wt./day. The pigs had free access to water via nipple drinkingsystems. The experimental diets were administered for 28 days. All theexperimental procedures described followed established guidelines for the careand use of laboratory animals and were approved by the local veterinary office.

Sample Collection. After completion of the feeding period, the animalswere killed under a light anesthesia. Blood was collected into heparinizedpolyethylene tubes. Liver and thyroid gland were dissected and weighed.Plasma was obtained by centrifugation of the blood (1100g; 10 min). All thesamples were stored at �80°C pending analysis.

Total RNA Preparation and cDNA Synthesis. Total RNA from liver andthyroid tissue was isolated by TRIzol reagent (Invitrogen, Karlsruhe, Ger-many) following the manufacturer’s protocol, resuspended in diethyl pyrocar-bonate-treated water, and stored at �80°C until use. The concentration of totalRNA was determined by ultraviolet absorbance at 260 nm. The quality of allthe RNA samples was assessed by agarose gel electrophoresis. cDNA wasprepared from total RNA (1.2 �g) by reverse transcription using M-MuLVreverse transcriptase (MBI Fermentas, St. Leon-Rot, Germany) and oli-go(dT)18 primers (Operon, Cologne, Germany).

Semiquantitative Polymerase Chain Reaction. Expression analysis forsemiquantitative reverse transcriptase-polymerase chain reaction (RT-PCR)was normalized using glyceraldehyde 3-phosphate dehydrogenase (GAPDH)as an internal standard. cDNA templates (2 �l) were used in a final volume of20 �l containing 0.2 �M concentration of the corresponding primers (Roth,Karlsruhe, Germany) (see Table 1), 1.5 mM magnesium chloride, 1� PCRbuffer, 1 U Taq polymerase (Gene Craft, Luedinghausen, Germany), and 0.2mM deoxyribonucleoside triphosphates (Roth). Each PCR cycle compriseddenaturation for 30 s at 94°C, annealing for 30 s at 60 to 64°C (see Table 1),and elongation for 1 min at 72°C, followed by a final extension period for 10min at 72°C. Number of cycles for each primer pair was tested previously.Cycle number was as follows: genes of thyroid gland: GAPDH, 23; sodiumiodide symporter, 25; dual oxidase 2, 42; cathepsin B, 35; TSH receptor, 28;type II iodothyronine deiodinase, 40; and thyroglobulin, 33; hepatic genes:GAPDH, 32; acyl CoA oxidase (ACO), 32; carnitine palmitoyl transferase 1(CPT-1), 32; albumin, 20; thyroid hormone receptor �1, 35; thyroxine-bindingglobulin, 42; transthyretin, 30; and type I iodothyronine deiodinase, 40. Awater control was included in all the PCRs for detection of contamination, anddilutions of the isolated total RNA corresponding to the cDNA synthesis wereused as template to exclude impurities caused by genomic DNA. A volume of10 �l per PCR was submitted to agarose gel electrophoresis (1.5%). Ethidiumbromide-stained gels were digitized for quantification (apparatus and softwarefrom Syngene, Cambridge, UK).

Preparation of Hepatic Microsomes. One gram of liver was homogenizedin a medium (10 ml) containing 0.25 M sucrose and 0.1 M phosphate buffer

TABLE 1

Sequences of primers used for semiquantitative RT-PCR

Gene (NCBI GenBank) Forward Primer Reverse Primer Size/AnnealingTemperature

bp/°C

Acyl CoA oxidase (AF185048) CTCGCAGACCCAGATGAAAT TCCAAGCCTCGAAGATGAGT 218/60Albumin (X12422) GCACGAGAAGACACCAGTGA CGAGTGCAGTTTGCTTCTTG 200/62CPT-I (AF288789) GCATTTGTCCCATCTTTCGT GCACTGGTCCTTCTGGGATA 198/60Cathepsin B (AJ315560) GGCCTCTATGACTCGCATGT GCAAGTTCCCCTCAAGTCTG 198/60Dual oxidase 2 (AF547267) GACCCAGCGGCAGTTTGAATGG AGGGCCGCAGCTGAACACTCC 295/64GAPDH (AF017079) AGGGGCTCTCCAGAACATCATCC TCGCGTGCTCTTGCTGGGGTTGG 446/60Sodium iodide symporter (AJ487855) AGTCATCAGCGGCCCCCTCCTC ACCGATGCCGTCTGCCGTGTG 456/60Thyroglobulin (AF165610) CAGTAAGGGCTTCCGTCTTG GGAGCTGCACTGAGGAATGT 198/60Thyroid hormone receptor �1 (AJ005797) CCAGATGGAAAGCGAAAAAG TGGGATGGAGATTCTTCTGG 199/60Thyroid peroxidase (X04645) CTGGGCGCCGTGCTCGTCTG ACGCGGGTGGCATCTGACTCTGAC 287/65Thyroxine-binding globulin (NM214058) GTGGCTTCTTGGGCATGTAT GAACCTCCGGTACAGGTTGA 206/62Transthyretin (X87846) ATGGTCAAAGTCCTGGATGC TGCCTTCCAGTAGGATTTGG 207/60TSH receptor (NM214297) GCCTGCCCATGGACACTGAGAC CTGACCCCGGTATGCCTGAGC 422/60Type I iodothyronine deiodinase (AY533206) CTCTGGGTGCTCTTTCAGGT ATCGGACCTTCAGCACAAAC 199/62Type II iodothyronine deiodinase (NM001001626) CTCGGTCATTCTCCTCAAGC TGCTTCCTTCAGGATTGGAG 200/60

1888 LUCI ET AL.

25

(pH 7.4) using a Potter-Elvehjem homogenizer. Homogenates were centri-fuged at 1000g for 10 min at 4°C, and the supernatant was centrifuged at15,000g for 15 min. The microsomal pellet was obtained by centrifugation ofthe 15,000g supernatant at 105,000g for 60 min. Microsomal pellets weresuspended in the homogenization medium, and the protein concentration wasdetermined with the bicinchoninic acid reagent according to the supplier’sprotocol using bovine serum albumin as standard.

Enzyme Assays. The activity of pNP-UGT in hepatic microsomes wasassayed by the method of Thurman et al. (1981). The assay mixture consistedof 50 mM potassium phosphate buffer (pH 7.4), 0.2 mg of bovine serumalbumin/ml, 1 mM magnesium chloride, 0.5 mM dithiothreitol, 0.5 mg ofTriton X-100/ml, 0.2 mM pNP, 0.2 mM UDPGA, and 0.25 mg of microsomalprotein/ml. The final volume of the assay was 200 �l. A blank was incubatedwithout UDPGA. The reaction was stopped after 30 min at 37°C by additionof 1 ml of 0.1 M sodium hydroxide solution. The pNP glucuronidation wasquantified by measuring the decrease of absorbance at 400 nm. The concen-tration of pNP glucuronide was calculated using a molar extinction coefficientof 18,300/cm.

Activity of bilirubin-UGT in hepatic microsomes was measured in an assaymixture containing 0.1 M Tris-hydrochloride (pH 7.8), 0.1 mM bilirubin, and5 mM UDPGA in a final volume of 200 �l. Suspended microsomes werepreincubated with 20 mg/ml sodium cholate (1:1, v/v) for 10 min at 4°C. Thereaction was started by addition of 1 mg of microsomal protein/ml for 60 minat 37°C. The amount of bilirubin glucuronide formed during the incubationwas quantified with a commercial kit (DiaSys Diagnostic Systems, Holzheim,Germany).

T3- and T4-UGT activities were determined in separate assays using amodified version of the method of Beetstra et al. (1991) by incubating 1 �MT3 or T4, respectively, and 0.1 �Ci of 125I-labeled T3 or T4 in reaction mixturecontaining 75 mM Tris-hydrochloride (pH 7.8), 7.5 mM magnesium chloride,0.25 mg of Brij 56/ml, 5 mM UDPGA, and 1 mM 6-propyl-2-thiouracil. Thefinal volume of the assay was 200 �l. Reactions were started by adding 0.5 mgof microsomal protein/ml at 37°C. Blanks were performed in the absence ofUDPGA. After 30 min, reactions were terminated by addition of 200 �l ofice-cold methanol, and the mixtures were centrifuged at 3500g for 8 min. Fiftymicroliters of the supernatants was injected into a high-performance liquidchromatograph for separation of T3 or T4 glucuronides formed during theincubation by a modified version of the method of Jemnitz et al. (2000). Thehigh-performance liquid chromatography equipment consisted of a 1100 seriespump (isocratic), an autosampler, a LiChrospher 100 RP 18e column (125 �4 mm, 5-�m particle size) with matching guard column (4 � 4 mm; AgilentTechnologies, Waldbronn, Germany). The mobile phase consisted of 50 mMpotassium dihydrogen phosphate and methanol (43:57, v/v, pH 7.0). Forseparation of T3-glucuronide, the flow rate was 0.8 ml/min. For separation ofT4-glucuronide, the flow rate was 1.25 ml/min. Fractions containing T3- orT4-glucuronide, respectively, were collected with a fraction collector 203(Gilson International, Bad Camberg, Germany). The radioactivity of the frac-tions was counted to calculate T3- and T4-UGT activities.

Histology of Thyroid Gland. Samples of thyroid glands were fixed byimmersion in 10% neutral buffered formalin, processed for embedding intoparaffin wax, and cut into 4-�m sections. For light microscopy, the sectionswere stained with hemalum and eosin. The epithelial cell height was measuredusing 4 cells per follicle in 100 follicles of each thyroid. The lumen diameterwas measured in 10 sections for 10 follicles per section of each thyroid. All thepictures were digitized, and the parameters were measured using the Lucia G(Nikon, Dusseldorf, Germany) software (release 4.81).

Analysis of Plasma Hormones. The plasma concentrations of free and totalT4 and total T3 were measured with radioimmunoassay kits (MP Biomedicals,Eschwege, Germany).

Statistics. The results were analyzed using Minitab (State College, PA)statistical software (release 13). Statistical significance of differences betweencontrol group and treatment group was evaluated using Student’s t test. Meanvalues were considered significantly different for P � 0.05.

Results

Initial and final body weights after an experimental period of 28days were similar in both groups of pigs (Table 2). Animals treated

with clofibrate had heavier livers (P � 0.05) and higher concentra-tions of microsomal protein in the liver (P � 0.05) than control pigs(Table 2). Relative hepatic mRNA concentration of the PPAR� targetgenes ACO and CPT-1 was higher (P � 0.05) in pigs treated withclofibrate than in control pigs (ACO: 1.39 � 0.27 versus 1.00 � 0.35;CPT-1: 1.60 � 0.13 versus 1.00 � 0.12; mean � S.D., n � 9 for eachgroup). Moreover, concentrations of total and free T4 and total T3 inplasma were markedly lower in pigs treated with clofibrate than incontrol pigs (P � 0.05), whereas the T4/T3 ratio did not differ betweenboth groups of pigs (Table 2).

Pigs treated with clofibrate had a higher activity of bilirubin-UGTin the liver than control pigs (1.08 � 0.05 versus 0.44 � 0.02nmol/min/mg; mean � S.D., n � 9 for each group; P � 0.05). Theactivity of hepatic pNP-UGT was lower in pigs treated with clofibratethan in control pigs (44 � 5 versus 70 � 8 nmol/min/mg; mean �S.D., n � 9 for each group; P � 0.05). Activities of hepatic T3- andT4-UGT were higher in pigs treated with clofibrate than in controlpigs (P � 0.05) (Fig. 1).

Weights of thyroids, diameter of follicle lumen, and thyroid epi-thelial cell height did not differ between both groups of pigs (Table 3).Relative mRNA concentrations of TSH receptor, sodium iodide sym-porter, thyroid peroxidase, and cathepsin B were higher in thyroids of

FIG. 1. Activities of T3- and T4-UGT in the liver of pigs fed a control diet or a dietsupplemented with 5 g of clofibrate/kg for 28 days. Data are reported as mean �S.D. with nine animals per group. �, significantly different to control group (P �0.05).

TABLE 3

Thyroid weight, follicle lumen diameter, and epithelial cell height in thyroid ofpigs fed a control diet or a diet supplemented with 5 g of clofibrate/kg for

28 days

Data are reported as mean � S.D. with nine animals per group.

Control Clofibrate

Thyroid weight (g) 2.17 � 0.19 2.39 � 0.55Follicle lumen diameter (�m) 84.9 � 10.9 88.8 � 14.2Epithelial cell height (�m) 7.53 � 0.14 7.64 � 0.67

TABLE 2

Body and liver weights, microsomal protein in the liver, and plasma thyroidhormone concentrations in pigs fed a control diet or a diet supplemented with

5 g of clofibrate/kg for 28 days

Data are reported as mean � S.D. with nine animals per group.

Control Clofibrate

Initial body weight (kg) 12.0 � 1.1 11.9 � 0.6Final body weight (kg) 26.0 � 1.5 25.2 � 1.2Liver weight (g) 673 � 63 779 � 63a

Hepatic microsomal proteins (mg/g liver) 12.4 � 1.8 15.4 � 2.4a

Plasma thyroid hormonesTotal thyroxine (T4, nM) 45.2 � 13.1 29.6 � 7.1a

Free thyroxine (pM) 12.6 � 3.7 8.6 � 2.2a

Total triiodothyronine (T3, nM) 1.20 � 0.52 0.64 � 0.11a

T4/T3 ratio 41.1 � 13.2 40.3 � 10.7

a P � 0.05 compared with control group.

1889CLOFIBRATE AND THYROID HORMONE GLUCURONIDATION IN PIGS

26

pigs treated with clofibrate than in thyroids of control pigs (P � 0.05)(Fig. 2). mRNA concentration of type II iodothyronine deiodinase waslower in thyroids of pigs treated with clofibrate than in thyroids ofcontrol pigs (P � 0.05); those of dual oxidase 2 and thyroglobulin didnot differ between both groups of pigs (Fig. 2).

Pigs treated with clofibrate had lower hepatic mRNA concentra-tions of transthyretin, albumin, and thyroid hormone receptor �1 thancontrol pigs (P � 0.05) (Table 4). Hepatic mRNA concentration ofthyroxine-binding globulin tended to be lower in pigs treated withclofibrate compared with control pigs (P � 0.10), whereas hepaticmRNA concentration of type I iodothyronine deiodinase did not differbetween both groups of pigs.

Discussion

To our knowledge, this is the first study to investigate the effect ofclofibrate on the hepatic metabolism of thyroid hormones in the pig.It is well known that pigs are a nonproliferating species, meaning thattreatment with PPAR� agonist causes no or only weak peroxisomeproliferation in the liver. Interestingly, in this study clofibrate treat-ment caused a significant increase in liver weights of pigs, by about15%. This is in disagreement with a recent study in which pigs did notshow significantly increased liver weights after a 1-week treatmentwith a dose of clofibrate similar to those used in our study (Cheon etal., 2005). The difference in these results could be because of thelonger treatment period in our study compared with that in the studyof Cheon et al. (2005). Moreover, we observed a moderate up-regulation of the PPAR� target genes ACO and CPT-1 in the liver ofpigs treated with clofibrate, which indicates that clofibrate treatmentcaused PPAR� activation in these pigs. Nevertheless, increases inliver weights and hepatic ACO and CPT-1 mRNA concentration weremuch lower than those observed in rodents treated with clofibrate. Inrats and mice, feeding PPAR� agonists increases liver weights by50% or more and mRNA concentrations of ACO 5- to 10-foldcompared with untreated controls (Kawashima et al., 1990; He et al.,2002; Frederiksen et al., 2004; Li et al., 2004).

Several families of UGT enzymes are expressed in the liver. Tostudy the effect of clofibrate treatment on the induction of microsomalenzymes, we determined the activities of bilirubin- and pNP-UGT.The finding that clofibrate treatment strongly increases bilirubin-UGTis in accordance with studies in rats and mice (Visser et al., 1993a;Viollon-Abadie et al., 1999). The finding that clofibrate reduces theactivity of pNP-UGT is also in accordance with a study in whichclofibrate significantly reduced the activity of pNP-UGT in the liverof Wistar rats (Visser et al., 1993a). These observations suggest that

clofibrate stimulated the microsomal enzyme system in pigs in asimilar way as in rats. This study also shows that clofibrate treatmentstrongly increases the activity of T3- and T4-UGT in the liver, whichin turn leads to a dramatic reduction of plasma T3 and T4 concentra-tions. Increased activity of T4-UGT had previously been observed inWistar rats but not in mice treated with clofibrate (Visser et al., 1993a;Viollon-Abadie et al., 1999). The increased activity of T3-UGT in pigstreated with clofibrate, however, is in strong contrast to rats, in whichclofibrate treatment did not increase T3-UGT activity (Visser et al.,1993a).

In rats, T4 is accepted as a substrate by hepatic bilirubin-UGT(UGT1A1) and phenol-UGT (UGT1A6), and it was shown that in-creased activities of these enzymes were associated with increasedglucuronidation of T4 in the liver (Beetstra et al., 1991; Magdalou etal., 1993; Visser et al., 1993a,b; Viollon-Abadie et al., 2000; Vanselland Klaassen, 2002). The enzymes involved in glucuronidation ofthyroid hormones in pigs have not yet been identified. The fact thatactivities of both bilirubin- and T4-UGT were increased suggests thatin pig liver T4 was also glucuronidated by bilirubin-UGT, as happensin rats. It is probable that enzymes other than UGT1A1 and UGT1A6can also be induced by clofibrate in rats and are involved in T4conjugation (Jemnitz et al., 2000). In rat liver, glucuronidation of T3,unlike glucuronidation of T4, is catalyzed by androsterone-UGT(Beetstra et al., 1991; Visser et al., 1993b). The increased T3-UGTactivity in pigs treated with clofibrate could therefore also have beencaused by an increased activity of androsterone-UGT, although thiswas not assayed in this study. The UGT in pig liver have been lessextensively investigated and have not yet been phenotyped. Therefore,it remains unknown which specific UGT were responsive for theincreased glucuronidation of T3 and T4 in pigs treated with clofibrate.

It has been shown that activation of PPAR� leads to transcriptionalup-regulation of the CYP4A genes, which are also constituents of themicrosomal biotransformation system in both proliferating and non-proliferating species (Lawrence et al., 2001; Cheon et al., 2005). It hasfurther been shown that some UGT isoforms (UGT1A9, UGT2B4) arePPAR� target genes (Barbier et al., 2003a,b). PPAR� is naturallyactivated during fasting, and Visser et al. (1996) showed that foodrestriction resulted in increased bilirubin and thyroid hormone UGTactivities in rats. These findings suggest that UGT catalyzing theglucuronidation of thyroid hormones may be transcriptionally up-regulated by activation of PPAR�. It is well known that expression ofPPAR� in the liver is much lower in nonproliferating species and thatthe response of many genes to PPAR� activation is weaker than inproliferating species. This is also true for up-regulation of microsomalCYP4A genes by treatment with PPAR� agonists (Lawrence et al.,2001; Cheon et al., 2005). If PPAR� plays a crucial role in the

FIG. 2. Relative mRNA concentrations of TSH receptor (TSHR), sodium iodidesymporter (NIS), dual oxidase 2 (DUOX2), thyroid peroxidase (TPO), thyroglob-ulin (TG), cathepsin B (Cat B), and type II iodothyronine deiodinase (ID-II) in thethyroid of pigs fed a control diet or a diet supplemented with 5 g of clofibrate/kg for28 days. All the mRNA concentrations were determined by semiquantitative RT-PCR and normalized by GAPDH. Data are reported as mean � S.D. with nineanimals per group. Data are expressed relative to mRNA concentrations of controlpigs (control � 1). �, significantly different to control group (P � 0.05).

TABLE 4

Relative hepatic mRNA concentrations of transthyretin, thyroxine-bindingglobulin, albumin, thyroid hormone receptor �1, and type I iodothyroninedeiodinase in pigs fed a control diet or a diet supplemented with 5 g of

clofibrate/kg for 28 days

Data are reported as mean � S.D. with nine animals per group. All the mRNAconcentrations were determined by semiquantitative RT-PCR and normalized by GAPDH.Data are expressed relative to mRNA concentrations of control pigs (control � 1).

Control Clofibrate

Transthyretin 1.00 � 0.14 0.87 � 0.12a

Thyroxine-binding globulin 1.00 � 0.14 0.89 � 0.18b

Albumin 1.00 � 0.11 0.87 � 0.11a

Thyroid hormone receptor �1 1.00 � 0.14 0.84 � 0.10a

Type I iodothyronine deiodinase 1.00 � 0.07 0.98 � 0.07

a P � 0.05 compared with control group.b P � 0.10 compared with control group.

1890 LUCI ET AL.

27

activation of UGT catalyzing thyroid hormone glucuronidation, theeffect of clofibrate on up-regulation of these enzymes in pigs wouldbe expected to be much lower than in proliferating species such as ratsor mice. But activation of T3- and T4-UGT by clofibrate was evenstronger in pigs than reported for rats or mice (Visser et al., 1993a;Viollon-Abadie et al., 1999). These findings suggest that activation ofPPAR� does not play a key role in clofibrate-induced up-regulation ofthyroid hormone UGT. Nevertheless, the role of PPAR� in the reg-ulation of thyroid hormone glucuronidation should be further inves-tigated.

The increased activities of T3- and T4-UGT make it highly probablethat the markedly reduced plasma concentrations of T3 and T4 in pigstreated with clofibrate are mainly caused by increased glucuronidationof these hormones in the liver. Because most T3 is generated inperipheral tissues, mainly the liver, by deiodination of T4, a reducedT3 concentration could potentially be caused by an inhibition of typeI iodothyronine deiodinase. Indeed, in the study of Visser et al.(1993a), clofibrate treatment of rats reduced the activity of thatenzyme, which might be responsible for the reduced concentration ofT3 observed in their study. We did not determine the activity but onlythe mRNA concentration of that enzyme in the liver, which was notinfluenced by clofibrate treatment. Interestingly, in contrast to hepatictype I iodothyronine deiodinase, type II deiodinase in the thyroidshowed a reduced mRNA concentration in pigs treated with clofibratecompared with control pigs. A reduced activity of type II deiodinase,which converts T4 to T3 in the thyroid, may play some role in thereduced T3 concentration in plasma. However, because the thyroidproduces less than 20% of total T3 (Findlay et al., 2000), a reducedactivity of type II deiodinase most probably plays a minor role in thereduced plasma concentration of T3. The reduction of plasma concen-trations of total T3 (by 47% versus control), free T4 (by 32% versuscontrol), and total T4 (by 35%) concentrations by clofibrate are alsostronger than those observed in Wistar rats. In Wistar rats, a dose of800 mg of clofibrate/kg b.wt./day reduced plasma T3 concentration by27% but did not reduce plasma total and free T4 (Visser et al., 1993a).In mice, a dose of 300 mg of clofibrate/kg b.wt./day reduced plasmafree T4 concentration by 13% but did not significantly reduce plasmaconcentration of free T3 (Viollon-Abadie et al., 1999). It is clear thatdifferent studies cannot be directly compared with each other, butthese data nevertheless suggest that pigs could be even more sensitiveto disruptions of thyroid hormone metabolism by clofibrate thanrodents.

Reduced plasma concentrations of T3 and T4 are expected toincrease the release of TSH from the pituitary gland. It has indeedbeen shown that microsomal enzyme inducers elevate TSH plasmaconcentrations in rodents, which in turn stimulates proliferation ofepithelial cells in thyroid tissue as a result of increased glucuronida-tion of thyroid hormones (e.g., Curran and DeGroot, 1991; De Sandroet al., 1991; Saito et al., 1991; Liu et al., 1995). As no assay wasavailable for measuring TSH concentration in plasma of the pigs, wedetermined mRNA concentrations of various genes in the thyroid thatare responsive to TSH treatment. The finding that mRNA concentra-tions of TSH receptor, sodium iodide symporter, thyroid peroxidase,and cathepsin B, all genes responsive to TSH, were moderatelyincreased by 40 to 70% suggests that the thyroid was stimulated by theincreased plasma concentration of TSH. This suggestion is confirmedby a study that showed that TSH plasma concentrations are increasedby microsomal enzyme inducers, which stimulate the glucuronidationof T3 (Klaassen and Hood, 2001). Our study further shows thatexpression levels of dual oxidase 2, a hydrogen peroxide-generatingsystem that constitutes the rate-limiting step of thyroid hormone

synthesis, and of thyroglobulin, a protein involved in thyroid hormonesynthesis and storage, are not altered by clofibrate treatment.

The finding that thyroid weights, epithelial cells, and follicle lumendiameter were not increased by clofibrate was unexpected and sug-gests that stimulation of the thyroid was moderate, only increasinggene expression of TSH-responsive genes in the thyroid, whereashistological alterations (i.e., increased epithelial cell height) may takelonger than 4 weeks to become evident.

The action of thyroid hormones like T3 is mediated by thyroidhormone receptors that belong to the family of nuclear hormonereceptors. The present study shows that clofibrate treatment reducesgene expression of thyroid hormone receptor �1 in the liver of rats.This finding agrees with a recent study in which bezafibrate down-regulated thyroid hormone receptors in rat liver (Bonilla et al., 2001).That study further showed that down-regulation of thyroid hormonereceptors was caused by activation of PPAR�. Therefore, it is likelythat in our study down-regulation of thyroid hormone receptor �1 inpigs treated with PPAR� was also caused by PPAR� activation byclofibrate. Down-regulation of thyroid hormone receptor implies thatthe biological activity of T3 may have been reduced in pigs treatedwith clofibrate.

Thyroxine-binding globulin, transthyretin, and albumin are the ma-jor plasma transport proteins in pigs (Janssen et al., 2002). Theseproteins are synthesized in the liver. We found in our study that geneexpression of these proteins in the liver was reduced by clofibratetreatment of pigs. In studies by Motojima et al. (1992, 1997), the sameeffect of clofibrate on expression of transthyretin was observed in ratsand several mouse strains, whereas there was no effect in PPAR�-nullmice. This suggests that down-regulation of transthyretin expressionwas induced by PPAR� activation. Consequently, down-regulation oftransthyretin and possibly also of thyroxine-binding globulin andalbumin could be the result of PPAR� activation by clofibrate. Ourdata suggest, although we did not measure concentrations of theseproteins in blood, that clofibrate treatment lowers not only concen-trations of thyroid hormones in plasma but also could reduce thetransport capacity for thyroid hormones.

In conclusion, this study shows for the first time that clofibratetreatment induces a strong activation of T3- and T4-UGT in pigs,leading to increased glucuronidation and markedly reduced plasmaconcentrations of these hormones. These alterations were accompa-nied by moderately increased mRNA concentrations of various TSH-responsive enzymes in the thyroid gland, reduced hepatic mRNAconcentrations of proteins involved in thyroid hormone transport, andthyroid hormone receptors. Because the pig represents a species thatdoes not respond with peroxisome proliferation to treatment withPPAR� agonists, the study shows that clofibrate treatment also dis-rupts the metabolism of thyroid hormones in nonproliferating species.

References

Barbier O, Duran-Sandoval D, Pineda-Torra I, Kosykh V, Fruchart JC, and Staels B (2003a)Peroxisome proliferator-activated receptor alpha induces hepatic expression of the human bileacid glucuronidating UDP-glucuronosyltransferase 2B4 enzyme. J Biol Chem 278:32852–32860.

Barbier O, Villeneuve L, Bocher V, Fontaine C, Torra IP, Duhem C, Kosykh V, Fruchart JC,Guillemette C, and Staels B (2003b) The UDP-glucuronosyltransferase 1A9 enzyme is aperoxisome proliferator-activated receptor alpha and gamma target gene. J Biol Chem 278:13975–13983.

Barter RA and Klaassen CD (1992a) Rat liver microsomal UDP-glucuronyltransferase activitytoward thyroxine: characterization, induction and form specificity. Toxicol Appl Pharmacol115:261–267.

Barter RA and Klaassen CD (1994) Reduction of thyroid hormone levels and alteration of thyroidfunction by four representative UDP-glucuronosyltransferase inducers in rats. Toxicol ApplPharmacol 128:9–17.

Barter RA and Klaassen CD (1992b) UDP-glucuronosyltransferase inducers reduce thyroidhormone levels in rats by an extrathyroidal mechanism. Toxicol Appl Pharmacol 113:36–42.

Beetstra JB, van Engelen JG, Karels P, van der Hoek HJ, de Jong M, Docter R, Krenning EP,Hennemann G, Brouwer A, and Visser TJ (1991) Thyroxine and 3,3�,5-triiodothyronine are

1891CLOFIBRATE AND THYROID HORMONE GLUCURONIDATION IN PIGS

28

glucuronidated in rat liver by different uridine diphosphate-glucuronyltransferases. Endocri-nology 128:741–746.

Bonilla S, Redonnet A, Noel-Suberville C, Groubet R, Pallet V, and Higueret P (2001) Effect ofa pharmacological activation of PPAR on the expression of RAR and TR in rat liver. J PhysiolBiochem 57:1–8.

Cheon Y, Nara TY, Band MR, Beever JE, Wallig MA, and Nakamura MT (2005) Induction ofoverlapping genes by fasting and a peroxisome proliferator in pigs: evidence of functionalPPAR alpha in nonproliferating species. Am J Physiol Regul Integr Comp Physiol 288:R1525–R1535.

Curran PG and DeGroot LJ (1991) The effect of hepatic enzyme-inducing drugs on thyroidhormones and the thyroid gland. Endocr Rev 12:135–150.

De Sandro V, Chevrier M, Boddaert A, Melcion C, Cordier A, and Richert L (1991) Comparisonof the effects of propylthiouracil, amiodarone, diphenylhydantoin, phenobarbital, and 3-meth-ylcholanthrene on hepatic and renal T4 metabolism and thyroid gland function in rats. ToxicolAppl Pharmacol 111:263–278.

Findlay KA, Kaptein E, Visser TJ, and Burchell B (2000) Characterization of the uridinediphosphate-glucuronosyltransferase-catalyzing thyroid hormone glucuronidation in man.J Clin Endocrinol Metab 85:2879–2883.

Frederiksen KS, Wulff EM, Sauerberg P, Mogensen JP, Jeppesen L, and Fleckner J (2004)Prediction of PPAR-alpha ligand-mediated physiological changes using gene expressionprofiles. J Lipid Res 45:592–601.

He WS, Nara TY, and Nakamura MT (2002) Delayed induction of delta-6 and delta-5 desatu-rases by a peroxisome proliferators. Biochem Biophys Res Commun 299:832–838.

Holden PR and Tugwood JD (1999) Peroxisome proliferator-activated receptor alpha: role inrodent liver cancer and species differences. J Mol Endocrinol 22:1–8.

Janssen OE, Lahner H, Grasberger H, Spring SA, Saller B, Mann K, Refetoff S, and EinspanierR (2002) Characterization and primary structures of bovine and porcine thyroxine-bindingglobulin. Mol Cell Endocrinol 186:27–35.

Jemnitz K, Veres Z, Monostory K, and Vereczkey L (2000) Glucuronidation of thyroxine inprimary monolayer cultures of rat hepatocytes: in vitro induction of UDP-glucuronosyltrans-ferases by methylcholanthrene, clofibrate, and dexamethasone alone and in combination. DrugMetab Dispos 28:34–37.

Kawashima Y, Musoh K, and Kozuka H (1990) Peroxisome proliferators enhance linoleic acidmetabolism in rat liver. Increased biosynthesis of omega 6 polyunsaturated fatty acids. J BiolChem 265:9170–9175.

Klaassen CD and Hood AM (2001) Effects of microsomal enzyme inducers on thyroid follicularcell proliferation and thyroid hormone metabolism. Toxicol Pathol 29:34–40.

Lawrence JW, Li Y, Chen S, DeLuca JG, Berger JP, Umbenhauer DR, Moller DE, and Zhou G(2001) Differential gene regulation in human versus rodent hepatocytes by peroxisomeproliferator-activated receptor (PPAR) �. J Biol Chem 34:31521–31527.

Li Y, Nara TY, and Nakamura MT (2004) Regulation of highly unsaturated fatty acid synthesis:a new physiological role of peroxisome proliferators-activated receptor �. FASEB J 18:A863.

Liu J, Liu Y, Barter RA, and Klaassen CD (1995) Alteration of thyroid homeostasis byUDP-glucuronosyltransferase inducers in rats: a dose-response study. J Pharmacol Exp Ther273:977–985.

Mackenzie PI, Owens IS, Burchell B, Bock KW, Bairoch A, Belanger A, Fournel-Gigleux S,Green M, Hum DW, Iyanagi T, et al. (1997) The UDP glycosyltransferase gene superfamily:recommended nomenclature update based on evolutionary divergence. Pharmacogenetics7:255–269.

Magdalou J, Fournel-Gigleux S, Pritchard M, and Siest G (1993) Peroxisome proliferators asinducers and substrates of UDP-glucuronosyltransferases. Biol Cell 77:13–16.

Mandard S, Muller M, and Kersten S (2004) Peroxisome proliferator-activated receptor � targetgenes. Cell Mol Life Sci 61:393–416.

Miners JO and Mackenzie PI (1991) Drug glucuronidation in humans. Pharmacol Ther 51:347–369.

Motojima K, Goto S, and Imanaka T (1992) Specific repression of transthyretin gene expressionin rat liver by a peroxisome proliferator clofibrate. Biochem Biophys Res Commun 188:799–806.

Motojima K, Peters JM, and Gonzalez FJ (1997) PPAR alpha mediates peroxisome proliferator-induced transcriptional repression of nonperoxisomal gene expression in mouse. BiochemBiophys Res Commun 230:155–158.

National Research Council (1998) Nutrient Requirement of Swine, 10th ed revised, NationalAcademy of Sciences, Washington DC.

Peters JM, Cheung C, and Gonzalez FJ (2005) Peroxisome proliferator-activated receptor-alphaand liver cancer: where do we stand? J Mol Med 83:774–785.

Rushmore TH and Kong AN (2002) Pharmacogenomics, regulation and signaling pathways ofphase I and II drug metabolizing enzymes. Curr Drug Metab 3:481–490.

Saito K, Kaneko H, Sato K, Yoshitake A, and Yamada H (1991) Hepatic UDP-glucuronyltransferase(s) activity toward thyroid hormones in rats: induction and effects onserum thyroid hormone levels following treatment with various enzyme inducers. Toxicol ApplPharmacol 111:99–106.

Thurman RG, Reinke LA, Belinsky S, Evans RK, and Kauffman FC (1981) Co-regulation of themixed-function oxidation of p-nitroanisole and glucuronidation of p-nitrophenol in the per-fused rat liver by carbohydrate reserves. Arch Biochem Biophys 209:137–142.

Vansell NR and Klaassen CD (2002) Increase in rat liver UDP-glucuronyltransferase mRNA bymicrosomal enzyme inducers that enhance thyroid hormone glucuronidation. Drug MetabDispos 30:240–246.

Viollon-Abadie C, Bigot-Lasserre D, Nicod L, Carmichael N, and Richert L (2000) Effects ofmodel inducers on thyroxine UDP-glucuronosyl-transferase activity in vitro in rat and mousehepatocyte cultures. Toxicol In Vitro 14:505–512.

Viollon-Abadie C, Lassere D, Debruyne E, Nicod L, Carmichael N, and Richert L (1999)Phenobarbital, �-naphtoflavone, clofibrate and pregnenolone-16�-carbonitrile do not affecthepatic thyroid hormone UDP-glucuronosyl transferase activity, and thyroid gland function inmice. Toxicol Appl Pharmacol 155:1–12.

Visser TJ, Kaptain E, van der Toor H, van Raaij JA, van den Berg KJ, Joe CT, van Engelen JG,and Brouwer A (1993a) Glucuronidation of thyroid hormone in rat liver: effects of in vivotreatment with microsomal enzyme inducers and in vitro assay conditions. Endocrinology133:2177–2187.

Visser TJ, Kaptain E, van Raaij JA, Joe CT, Ebner T, and Burchell B (1993b) MultipleUDP-glucuronyltransferases for the glucuronidation of thyroid hormone with preference for3,3�,5�-triiodothyronine (reverse T3). FEBS Lett 315:65–68.

Visser TJ, van Haasteren GA, Linkels E, Kaptein E, van Toor H, and de Greef WJ (1996)Gender-specific changes in thyroid hormone-glucuronidating enzymes in rat liver duringshort-term fasting and long-term food restriction. Eur J Endocrinol 135:489–497.

Zhou J, Zhang J, and Xie W (2005) Xenobiotic nuclear receptor-mediated regulation ofUDP-glucuronosyl-transferases. Curr Drug Metab 6:289–298.

Address correspondence to: Klaus Eder, Institut fur Ernahrungswissen-schaften, Martin-Luther-Universitat Halle-Wittenberg, Emil-Abderhalden-Str. 26,D-06108 Halle/S., Germany. E-mail: [email protected]

1892 LUCI ET AL.

29

BioMed Central

Page 1 of 11(page number not for citation purposes)

BMC Pharmacology

Open AccessResearch articleClofibrate treatment in pigs: Effects on parameters critical with respect to peroxisome proliferator-induced hepatocarcinogenesis in rodentsSebastian Luci1, Beatrice Giemsa1, Gerd Hause2, Holger Kluge1 and Klaus Eder*1

Address: 1Institut für Agrar- und Ernährungswissenschaften, Martin-Luther-Universität Halle-Wittenberg, Emil-Abderhalden-Strasse 26, D-06108 Halle (Saale), Germany and 2Biozentrum, Martin-Luther-Universität Halle-Wittenberg, Weinbergweg 22, 06120 Halle (Saale), Germany

Email: Sebastian Luci - [email protected]; Beatrice Giemsa - [email protected]; Gerd Hause - [email protected]; Holger Kluge - [email protected]; Klaus Eder* - [email protected]

* Corresponding author

AbstractBackground: In rodents treatment with fibrates causes hepatocarcinogenesis, probably as a resultof oxidative stress and an impaired balance between apoptosis and cell proliferation in the liver.There is some debate whether fibrates could also induce liver cancer in species not responsive toperoxisome proliferation. In this study the effect of clofibrate treatment on peroxisomeproliferation, production of oxidative stress, gene expression of pro- and anti-apoptotic genes andproto-oncogenes was investigated in the liver of pigs, a non-proliferating species.

Results: Pigs treated with clofibrate had heavier livers (+16%), higher peroxisome counts (+61%),higher mRNA concentration of acyl-CoA oxidase (+66%), a higher activity of catalase (+41%) butlower concentrations of hydrogen peroxide (-32%) in the liver than control pigs (P < 0.05);concentrations of lipid peroxidation products (thiobarbituric acid-reactive substances, conjugateddienes) and total and reduced glutathione in the liver did not differ between both groups. Clofibratetreated pigs also had higher hepatic mRNA concentrations of bax and the proto-oncogenes c-mycand c-jun and a lower mRNA concentration of bcl-XL than control pigs (P < 0.05).

Conclusion: The data of this study show that clofibrate treatment induces moderate peroxisomeproliferation but does not cause oxidative stress in the liver of pigs. Gene expression analysisindicates that clofibrate treatment did not inhibit but rather stimulated apoptosis in the liver ofthese animals. It is also shown that clofibrate increases the expression of the proto-oncogenes c-myc and c-jun in the liver, an event which could be critical with respect to carcinogenesis. As theextent of peroxisome proliferation by clofibrate was similar to that observed in humans, the pigcan be regarded as a useful model for investigating the effects of peroxisome proliferators on liverfunction and hepatocarcinogenesis.

Published: 16 April 2007

BMC Pharmacology 2007, 7:6 doi:10.1186/1471-2210-7-6

Received: 10 January 2007Accepted: 16 April 2007

This article is available from: http://www.biomedcentral.com/1471-2210/7/6

© 2007 Luci et al; licensee BioMed Central Ltd. This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

30

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?cmd=Retrieve&db=PubMed&dopt=Abstract&list_uids=17437637

http://www.biomedcentral.com/1471-2210/7/6

http://creativecommons.org/licenses/by/2.0

http://www.biomedcentral.com/

http://www.biomedcentral.com/info/about/charter/

BMC Pharmacology 2007, 7:6 http://www.biomedcentral.com/1471-2210/7/6


BackgroundPeroxisome proliferators (PPs) comprise a diverse groupof chemicals, including pharmaceuticals, industrial chem-icals, endogenous fatty acids and eicosanoids. They bindto and activate the peroxisome proliferator-activatedreceptor (PPAR)-α, a transcription factor belonging to thenuclear hormone receptor superfamily [1]. Activation ofPPARα causes an increase in the transcription of genesrelated to fatty acid transport across the cell membrane,intracellular lipid trafficking, mitochondrial and peroxi-somal fatty acid uptake, and both mitochondrial and per-oxisomal fatty acid β-oxidation [2]. Administration of PPsto rats and mice typically causes hepatic peroxisome pro-liferation, hypertrophy, hyperplasia, and hepatocarcino-genesis [3,4]. PPARα-induced hepatocarcinogenesis inrats and mice may be mainly due to an increased oxidativestress caused by peroxisome proliferation and an altera-tion of the balance between apoptosis and cell prolifera-tion [5,6]. Treatment with PPs such as fibrates causes a 15to 20-fold up-regulation of acyl-CoA oxidase (ACO) andother peroxisomal oxidases that lead to the production ofhydrogen peroxide (H2O2) which under normal non-induced circumstances can be detoxified by catalase [7,8].Catalase induction increases only approximately twofoldin response to PPARα agonists in rodents, and the activityof glutathione peroxidase is often depressed followinglong term administration of PPs [9,10]. The capacity ofthe H2O2-degrading enzymes therefore may be insuffi-cient to detoxify the large increase in H2O2. Increased cel-lular H2O2 could also react with metals and generatehighly reactive hydroxyl radicals that could damage DNA,proteins or lipids [7]. Indeed, oxidatively damaged DNAand peroxide-modified lipids have been found in hepato-cytes of rats treated with PPs [11-13]. Activation of PPARαalso leads to increases in hepatocellular proliferation andinhibition of apoptosis, and when this occurs in DNA-damaged cells, it is thought to lead to proliferation of ini-tiated cells progressing to liver tumour [14,15]. ThatPPARα is required to mediate hepatocarcinogenesis byPPs has been demonstrated in studies with PPARα-nullmice that are refractory to this in response to long termadministration of PPs [15,16].

It is well known that non-human primates and humansare only weakly responsive to peroxisome proliferation incomparison to rodents [17,18]. Nevertheless, there is con-siderable controversy as to whether the administration ofdrugs which are ligands for PPARα to humans causes livercancer. This is significant because PPARα agonists such asfibrates have been in clinical use for the treatment ofhyperlipidaemias for many years. The data regarding theability of fibrates to cause peroxisome proliferation inhumans are diverse. Examination of liver biopsy samplesfrom patients receiving therapeutic doses of PPARα ago-nists showed a slight increase in peroxisome counts [19],

while others showed no increase [20,21]. In non-humanprimates, administration of clofibrate induced a moder-ate, dose-dependent peroxisome proliferation [22-24]. Itis therefore conceivable that cellular events induced byPPs related to hepatocarcinogenesis in rodents couldoccur also in non-proliferating species, albeit probablyless pronounced than in rodents.

Pigs, like humans and non-human primates, are a non-proliferating species [25]. They may therefore be a valua-ble model for investigating the effects of PPs on peroxi-some proliferation and related processes. To ourknowledge, the effect of fibrates on parameters related tohepatocarcinogenesis has not yet been investigated inpigs. We therefore treated pigs with clofibrate and deter-mined hepatic weight, number of peroxisomes and ACOexpression to provide information about the potency offibrates to induce peroxisome proliferation in pigs. Wealso considered the antioxidant status of the pigs (mRNAconcentrations and activities of various antioxidantenzymes including catalase, generation of H2O2 in theliver, concentrations of lipid peroxidation products) tofind out whether clofibrate causes oxidative stress in pigliver. In order to ascertain whether clofibrate treatmentcould affect the balance between cell proliferation andapoptosis we determined mRNA concentrations of pro-and anti-apoptotic genes, namely bax, bcl-XL and p53tumour suppressor gene. It has been shown that thenuclear factor κB (NF-κB) pathway is important in theactivation of genes that regulate cell proliferation andapoptosis in various cell types [26,27]. It was shownrecently that NF-κB contributes to the proliferative andapoptotic changes that occur in liver in response tofibrates [28]. More recently, it has been demonstrated thatthe p50 subunit of the NF-KB family is necessary for thepromotion of hepatocarcinogenesis by PPs [29]. To findout whether clofibrate treatment activated NF-KB in pigliver, we determined the mRNA concentration of tumornecrosis factor α (TNFα), a target gene of NF-KB which hasalso been identified as a suppressor of apoptosis and aninducer of DNA synthesis [30,31]. In rat liver and inmouse liver epithelial cells, treatment with the PPARαagonist WY-14,643 strongly up-regulated gene expressionof various proto-oncogenes including c-fos, c-jun and c-myc [32-35]. In mouse liver cells these changes were fol-lowed by enhanced DNA synthesis, and it has been con-cluded that this could play an important role in tumourpromotion by PPs [33]. Whether PPs stimulate expressionof proto-oncogenes in pigs has not yet been investigated.We therefore also determined gene expression of c-myc, c-jun and c-fos in liver of pigs treated with clofibrate, whichcould be critical with respect to hepatocarcinogenesis.

31



ResultsDue to the controlled feeding system, diet intake duringthe whole experimental period was identical in bothgroups of pigs, being 696 ± 7 g/d in average of the wholeperiod. Final body weights of the pigs on day 29 did notdiffer between the control group and the group treatedwith clofibrate (26.0 ± 1.5 kg for control pigs vs. 25.2 ± 1.2kg for pigs treated with clofibrate, n = 9 in each group).Pigs treated with clofibrate had higher liver weights(+16% in absolute terms, +19% in relative terms,expressed per kg body weight), higher peroxisome countsin the liver (+61%) (P < 0.05, Table 1). Relative mRNAconcentration of PPARα in liver did not differ betweenboth groups of pigs (control: 1.00 ± 0.38; clofibrate: 0.92± 0.20, n = 9, means ± SD) but pigs treated with clofibratehad a higher relative mRNA concentration of ACO in theliver (+66%) than control pigs (P < 0.05, Table 1).

As reference values for the expression of enzyme activities,we determined concentrations of protein in liver homoge-nate and liver cytosol. Protein concentration in liverhomogenate did not differ between both groups of pigs(control: 20.8 ± 5.9 mg/g liver; clofibrate: 21.1 ± 5.8 mg/g liver, n = 9, means ± SD); protein concentration in livercytosol was higher in pigs treated with clofibrate than incontrol pigs (control: 27.2 ± 2.3 mg/g liver; clofibrate:30.4 ± 3.0 mg/g liver, n = 9, means ± SD, P < 0.05). Pigstreated with clofibrate had higher mRNA concentrationsand activities of superoxide dismutase (SOD) (+77% and+128%, respectively) and catalase (+72% and +41%,respectively) in the liver than control pigs (P < 0.05, Table2). In contrast, mRNA concentration and activity of glu-tathione peroxidase (GSH-Px) in liver were reduced by26% and 15%, respectively, in pigs treated with clofibratecompared to control pigs (P < 0.05, Table 2). mRNA con-centration and activity of glutathione S-transferase (GST)in liver cytosol and concentrations of total and reducedglutathione in liver homogenate did not differ betweenboth groups of pigs (Table 2). However, the concentrationof α-tocopherol, both in absolute terms and in relativeterms, expressed per mmol of triglycerides + total choles-terol, was lower in the liver of pigs treated with clofibratethan in the liver of control pigs (-40%, P < 0.05) (Table 2).

The concentration of H2O2 in the liver was about 32%lower in pigs treated with clofibrate than in control pigs (P< 0.05, Table 3). Concentrations of lipid peroxidationproducts, thiobarbituric acid-reactive substances (TBARS)and conjugated dienes did not differ between both groupsof pigs, both in absolute terms and in relative terms,expressed per mmol of triglycerides + total cholesterol(Table 3).

Hepatic mRNA concentrations of p53 and c-fos did notdiffer between pigs treated with clofibrate and control pigs

whereas mRNA concentrations of bax, c-jun and c-mycwere higher in pigs treated with clofibrate than in controlpigs (P < 0.05, Fig. 1). Hepatic mRNA concentrations ofbcl-XL and TNFα were lower in pigs treated with clofibratethan in control pigs (P < 0.05, Fig. 1).

DiscussionTo our knowledge, this is the first study to investigate theeffect of clofibrate treatment on peroxisome proliferationand parameters that may be related to hepatocarcinogen-esis in pigs, a non-proliferating species. As in many otherstudies dealing with the effects of clofibrate on metabo-lism in experimental animals, we added clofibrate to thediet. The concentration of clofibrate in the diet of 5 g perkg diet was adopted from other studies with pigs[25,36,37]. The resulting daily dose of 220 mg per kgbody weight was relatively high compared with dosesused in humans for treatment of hyperlipidaemia, whichare usually in the range between 25 and 30 mg per kgbody weight.

Analysis of liver weights and number of peroxisomesshowed that treatment with clofibrate caused moderateperoxisome proliferation in pigs. The increase in thenumber of peroxisomes (+62%) observed in pigs treatedwith clofibrate is of a similar order of magnitude as the50% increase in liver peroxisome counts observed inhumans treated with clofibrate [19]. The extent of thechange in peroxisome counts in pigs is modest when com-pared with that reported for rodents. In rodents, adminis-tration of a dose of 200 mg clofibrate per kg body weight,comparable with that used in this study in pigs, resultedin a three- to five-fold increase in peroxisome counts com-pared with controls [38]. The current study also showsthat gene expression of ACO is moderately increased byclofibrate in pigs, which is in close accord with the mod-erate effect of clofibrate on the peroxisome count. Thepresent study confirms other studies [25,36,37] whichhave also shown that clofibrate causes only a relativelyweak up-regulation of PPARα target genes in pig liver. Themoderate effect of clofibrate on ACO in pigs is in strongcontrast to rodents where treatment with PPs causes a 10to 20-fold increase in ACO expression [35,39]. Up-regula-tion of ACO in the liver is critical because it leads toincreased production of H2O2 which causes oxidativestress within the cell. In contrast to observations inrodents, pigs treated with clofibrate in this study had nota higher but a lower concentration of H2O2 in the liver.We assume that this is due to up-regulation of catalase, thekey enzyme involved in H2O2 detoxification. In our studyboth ACO and catalase were up-regulated by clofibrate toa similar extent. Catalase has a high H2O2-detoxifyingactivity and is the rate-limiting enzyme for inhibitingH2O2 leakage from peroxisomes [40]. The finding that theconcentration of H2O2 was reduced even though the activ-

32



ity of GSH-Px, another H2O2-detoxifying enzyme local-ised in cytosol was reduced as well, suggests that the up-regulation of catalase was sufficient to eliminate all of theH2O2 produced in peroxisomes. The finding of a reducedactivity of GSH-Px in liver of pigs treated with clofibrateagrees with findings in rodents treated with PPs whichalso showed a lower activity of that enzyme in the liver[11,41].

Unsaturated fatty acids are susceptible to reactive oxygenspecies and undergo oxidation. The determination oflipid peroxides such as TBARS or conjugated dienes istherefore a sensible method to detect oxidative stress.Indeed, in hepatocytes of rats treated with PPs, concentra-tions of lipid peroxidation products were increased due tooxidative stress induced by peroxisome proliferation[11,42]. The fact that concentrations of TBARS and conju-gated dienes were not increased in the liver of pigs treatedwith clofibrate indicates that the moderate peroxisomeproliferation was not accompanied by oxidative stress.This indication is supported by the observation that the

concentration of reduced glutathione in the liver was alsonot altered in pigs treated with clofibrate when comparedwith control pigs. Glutathione plays a pivotal role in pro-tecting cells against the noxious effects of oxidant agents,and oxidative stress leads to enhanced oxidation of glu-tathione, which in turn causes a lower concentration ofreduced glutathione and a lower ratio of reduced to oxi-dized glutathione [43]. Likewise, activity of GST, anenzyme belonging to the phase II enzymes whose reac-tions include the addition of glutathione to electrophilicmolecules as well as the detoxification of organichydroperoxides, was not changed in pigs treated with clof-ibrate. This is in contrast to rodents treated with fibrates orother PPs that have a strongly reduced activity of GST inthe liver [44,45]. As it has been suggested that the reducedGST activity in rodents treated with PPs is the conse-quence of oxidative stress due to peroxisome prolifera-tion, the finding of an unchanged activity of that enzymeis another indication that oxidative stress did not occur inpigs treated with clofibrate. This indication agrees with arecent study which investigated the effect of high doses of

Table 1: Liver weights, number of peroxisomes and relative acyl-CoA oxidase mRNA concentration in the liver of pigs fed a control diet or a diet supplemented with 5 g clofibrate per kg for 28 days

Treatment Control (n = 9) Clofibrate (n = 9)

Liver weight (g) 673 ± 63 779 ± 63*Liver weight (g/kg body weight) 25.9 ± 2.2 30.9 ± 2.6*Number of peroxisomes (n/1,000 print) 366 ± 67 590 ± 116*Acyl-CoA oxidase mRNA 1.00 ± 0.35 1.66 ± 0.41*

Results are means ± SD.*Significantly different from control group (P < 0.05)

Table 2: Relative mRNA concentrations and activities of antioxidant enzymes and concentrations of some antioxidants in the liver of pigs fed a control diet or a diet supplemented with 5 g clofibrate per kg for 28 days


CatalasemRNA concentration 1.00 ± 0.49 1.72 ± 0.58*Activity (U/mg protein) 0.75 ± 0.14 1.06 ± 0.16*

Glutathione peroxidasemRNA concentration 1.00 ± 0.17 0.74 ± 0.18*Activity (U/mg protein) 4.7 ± 0.8 4.0 ± 0.4*

Glutathione S-transferasemRNA concentration 1.00 ± 0.41 0.90 ± 0.32Activity (U/mg protein) 0.76 ± 0.26 0.76 ± 0.18

Superoxide dismutasemRNA concentration 1.00 ± 0.42 1.77 ± 0.40*Activity (U/mg protein) 43 ± 8 97 ± 13*

Glutathione, total (nmol/mg protein) 2.13 ± 0.51 1.97 ± 0.90Glutathione, reduced (nmol/mg protein) 1.70 ± 0.50 1.67 ± 1.02α-tocopherol

(nmol/g) 14.5 ± 2.5 8.7 ± 2.9*[nmol/(mmol triglycerides + cholesterol)] 2.32 ± 0.40 1.36 ± 0.45*

Results are means ± SD.*Significantly different from control group (P < 0.05)#Relative to control (= 1.00)

33



fenofibrate and ciprofibrate in cynomolgus monkeys [24].In that study, clofibrate treatment induced moderatehepatic peroxisome proliferation, similar to that observedin the pigs of the present study, but there was also mini-

mal indication of oxidative stress. It is therefore con-cluded that even high doses of fibrates cause littleoxidative stress in the liver of non-proliferating species.

Relative mRNA concentrations of pro- and anti-apoptotic genes (bax, bcl-XL, p53, TNFα) and proto-oncogenes (c-myc, c-jun, c-fos) in livers of pigs fed a control diet or a diet supplemented with 5 g clofibrate per kg for 28 daysFigure 1Relative mRNA concentrations of pro- and anti-apoptotic genes (bax, bcl-XL, p53, TNFα) and proto-oncogenes (c-myc, c-jun, c-fos) in livers of pigs fed a control diet or a diet supplemented with 5 g clofibrate per kg for 28 days. All mRNA concentrations were determined by real-time quantitative PCR and normalized to GAPDH. Data are reported as means ± SD with nine ani-mals per group. Data are expressed relative to mRNA concentrations of control pigs (control = 1). * Significantly different to control group (P < 0.05).

Table 3: Concentration of hydrogen peroxide and lipid peroxidation products in the liver of pigs fed a control diet or a diet supplemented with 5 g clofibrate per kg for 28 days


Hydrogen peroxide (fluorescence/g liver) 29,437 ± 8,361 20,078 ± 7,225*Thiobarbituric acid-reactive substances

(nmol/g liver) 7.2 ± 1.6 7.7 ± 2.7[nmol/(mmol triglycerides + cholesterol)] 1.15 ± 0.26 1.20 ± 0.42

Conjugated dienes (nmol/g)(nmol/g liver) 16 ± 2 16 ± 2[nmol/(mmol triglycerides + cholesterol)] 2.56 ± 0.32 2.49 ± 0.31

Results are means ± SD.*Significantly different from control group (P < 0.05)

34



In the current study, pigs treated with clofibrate hadincreased hepatic mRNA concentration and activity ofSOD, an enzyme that converts superoxide anions intoH2O2 and therefore contributes to increased H2O2 produc-tion. The finding that H2O2 concentration in the liver wasreduced in spite of the increased activity of SOD is anotherindication that pigs treated with fibrate had a high hepaticH2O2-detoxifying capacity. Results from other studiesdealing with the effects of PPs on hepatic SOD are contra-dictory. In some studies, treatment of rodents with PPslowered hepatic activity of SOD [41,46]; in others treat-ment with PPs increased the activity of hepatic SOD[47,48]. Moreover, it has been shown that different PPscan have different effects on hepatic activity of SOD, andthat there are also differences between mice and hamstersin the effect of PPs on SOD activity [48]. The reason forthese contradictory results is unclear and should be inves-tigated further. As SOD is an important constituent of thehepatic antioxidant system, an increased activity of thatenzyme observed in pigs treated with clofibrate couldcontribute to a high antioxidant capacity in the liver ofthese pigs.

The finding that pigs treated with clofibrate had a reducedconcentration of α-tocopherol in the liver agrees withthose of several other studies in which rodents weretreated with PPs [11,48,49]. It has been shown that thetocopherol-lowering effect of PPs is stronger in rats thanin hamsters, suggesting that there is a correlation with thedegree of peroxisome proliferation [48]. It has been sug-gested that the reduction of hepatic tocopherol concentra-tion is not primarily due to oxidative stress produced byPPs but rather to their hypolipidaemic effect [48]. Toco-pherols are transported by lipoproteins within the bodyand as hypolipidaemic drugs reduce the number of lipo-protein particles in blood, they could also impair vitaminE transport in the body, i.e. transport of vitamin E fromthe intestine to the liver by chylomicrons. The fact that α-tocopherol concentration was reduced in the liver of pigstreated with clofibrate although there were no signs of oxi-dative stress confirms the suggestion that PPs do reducehepatic vitamin E concentrations independent of oxida-tive stress.

It has been shown that peroxisome proliferation leads tohepatocellular proliferation and to inhibition of apopto-sis, and these processes may contribute to hepatomegalyand to hepatocarcinogenesis observed in rodents treatedwith PPs [14,15]. The present study shows that clofibratetreatment of pigs increases gene expression of the proto-oncogenes c-jun and c-myc, which are required for entryinto the S phase of the cell cycle. These findings are inagreement with recent studies in rodent livers and livercells in which treatment with WY-14,643 strongly up-reg-ulated gene expression of proto-oncogenes [32-34]. Up-

regulation of proto-oncogenes in mouse liver cells wasfollowed by enhanced progression, and it has been sug-gested that this could play a role in tumour promotion ofPPs [32]. Although up-regulation of proto-oncogenes wassmaller in pigs treated with fibrates than in rodents [32-34], increased levels of c-jun and c-myc could haveenhanced cell proliferation, which might explain theincreased liver mass in pigs treated with clofibrate. Up-regulation of proto-oncogenes could be a critical eventwith respect to tumorigenesis. On the other hand, c-myccan collaborate with other proteins to induce apoptosisand sensitize cells to a variety of apoptotic triggers[50,51]. In order to find out whether clofibrate treatmentin pigs could have altered apoptosis, we determinedmRNA concentrations of bax, bcl-XL and p53 in the liver.Genes of the bcl-2 family have anti-apoptotic effects,which is antagonized by bax. The bcl-2/bax ratio is a keyfactor for determining apoptosis. When bcl-2 is expressedexcessively, bcl-2-bax heterogenous dimer predominates,thus inhibiting apoptosis [52]. When bax is expressedexcessively, bax-bax homogenous dimer or monomer pre-dominates, thus promoting apoptosis. p53 is a tumoursuppressor which exerts control over cell cycling by con-trolling the progression through the G1 phase [53]. Gen-otoxic stress or DNA damage leads to nuclearaccumulation of p53, which in turn activates the tran-scription of several genes involved in DNA repair or apop-tosis, including bcl-2 [54]. We did not directly determineapoptosis in the liver of piglets but the finding that expres-sion of bax was up-regulated in pigs treated with clofibratewhile expression of bcl-XL was reduced, together with theobservation of unchanged p53 expression suggests thatapoptosis was not inhibited but could instead have beenincreased in these pigs. This suggestion agrees with recentstudies which have shown that clofibrate induces apopto-sis in human and rat hepatoma cells [55-59]. In agree-ment with our study, a recent study showed that treatmentwith WY 14,643 up-regulates pro-apoptotic genes anddown regulates anti-apoptotic genes in the liver of mice,an effect which did not occur in PPARα-null mice [60]. Inthat study, it was also demonstrated that PPARα activa-tion increases the sensitivity of liver towards apoptosis byJo-2, an inducer of hepatic apoptosis. Therefore, it hasbeen suggested that PPARα could serve as a pharmacolog-ical target in diseases where apoptosis is a contributingfeature [60,61].

In a recent study in rats it was found that NF-KB is acti-vated by WY-14,643, probably due to oxidative stresscaused by peroxisome proliferation [62]. It has been dem-onstrated that NF-KB is essential for inducing cell prolifer-ation and hepatocarcinogenesis in rodents treated withfibrates [28,29]. The finding that mRNA concentration ofTNFα, a target gene of NF-κB, was reduced in liver of pigstreated with clofibrate compared to control pigs indicates

35



that clofibrate did not activate the NF-KB pathway in theseanimals. This finding may be related to the observationthat clofibrate treatment did not cause oxidative stress inpigs, in contrast to rats. The finding that WY-14,643 didnot activate the NF-KB pathway in hamsters either [62]suggests that PPs activate NF-KB only in species that areresponsive to PP induced hepatocarcinogenesis.

ConclusionTreatment with clofibrate at doses higher than those usedfor hypolipidaemic treatment in humans causes moderateperoxisome proliferation in the liver of pigs. Determina-tion of the concentration of H2O2 and lipid peroxidationproducts indicates that this did not produce oxidativestress. Determination of mRNA concentrations of pro-and anti-apoptotic genes in the liver indicates that clofi-brate treatment did also not inhibit but rather stimulatedapoptosis in these animals. Up-regulation of the proto-oncogenes c-myc and c-jun in the liver, however, could bea critical event with respect to carcinogenesis, whichdeserves further investigation in future studies. As theextent of peroxisome proliferation by clofibrate was simi-lar to that observed in humans, the pig can be regarded asa useful model for the investigating the effects of PPs onliver function and hepatocarcinogenesis.

MethodsAnimals and treatmentsEighteen male 8-week-old crossbred pigs [(German Lan-drace × Large White) × Pietrain] were kept in a roomunder controlled conditions at 23 ± 2°C and 55 ± 5% rel-ative humidity with light from 0600 to 1800 h. One daybefore the start of the experimental feeding period the pigswere weighed and randomly assigned to two groups withbody weights of 12.0 ± 1.1 kg in the control group and11.9 ± 0.6 kg in the treatment group. Both groups of pigsreceived a nutritionally adequate dry diet for growing pigs(according to [63]) containing (in g/kg) wheat (400), soy-bean meal (230), wheat bran (150), barley (100), sun-flower oil (90) and mineral premix including L-lysine,DL-methionine and L-threonine (30). This diet contained14.4 MJ metabolizable energy and 185 g crude protein perkg. The whole daily amount of diet was administered onceat 8.00 h. Diet intake was controlled, and each animal inthe experiment was offered an identical amount of dietper day. The amount of diet administered was about 15%below that consumed ad-libitum by pigs of a similarweight (as assessed in a previous study). Therefore, thediet offered was completely taken up by all pigs in theexperiment. During feeding period, the amount of dietoffered each day was increased continuously from 400 to1,200 g. Pigs of both groups received the same diet. How-ever, pigs of the treatment group were given additionally5 g clofibrate per kg diet which was freshly given onto thediet on each day. The pigs had free access to water via nip-

ple drinking systems. The experimental diets were admin-istered for 28 d. All experimental procedures describedfollowed established guidelines for the care and use oflaboratory animals and were approved by the local veter-inary office.

Sample collectionAfter completion of the feeding period the piglets werecaptive-bolt stunned and exsanguinated. Four hoursbefore euthanasia each pig was fed its respective diet. Afterkilling, blood was collected into heparinized polyethyl-ene tubes. Plasma was obtained by centrifugation of theblood (1,100 × g; 10 min; 4°C). The liver was dissectedand weighed and samples were stored at -80°C until anal-ysis. For preparation of liver homogenate, one g of liverwas homogenized in 10 mL of 0.1 M phosphate buffer,pH 7.4, containing 0.25 M sucrose using a Potter-Elve-hjem homogenizer. Homogenates were centrifuged at1,000 × g for 10 min at 4°C and the supernatant was cen-trifuged at 15,000 × g for 15 min again. The supernatantof that centrifugation was collected and centrifuged at105,000 × g for 60 min to yield the cytosolic fraction.Liver homogenates and cytosolic fraction were stored at -20°C for further analysis. Protein concentrations of liverhomogenates and cytosol were determined with the bicin-choninic acid reagent according to the supplier's protocol(Interchim, Montelucon, France) using bovine serumalbumin as the standard.

RT-PCR analysisTotal RNA from liver tissue was isolated by TRIzol reagent(Invitrogen, Karlsruhe, Germany) following the manufac-turer's protocol, resuspended in diethyl pyrocarbonate-treated water and stored at -80°C until use. The concentra-tion and purity of total RNA was determined by ultravioletabsorbance at 260 and 280 nm (SpectraFluor Plus; Tecan,Crailsheim, Germany). The quality of all RNA sampleswas assessed by agarose gel electrophoresis. cDNA wasprepared from total RNA (1.2 μg) by reverse transcriptionusing M-MuLV Reverse Transcriptase (MBI Fermentas, St.Leon-Rot, Germany) and oligo(dT)18 primers (OperonBiotechnologies, Cologne, Germany). The mRNA concen-tration of genes was measured by realtime detection PCRusing SYBR® Green I and a MJ Research Opticon system(Biozym Diagnostik GmbH, Oldendorf, Germany). Real-time detection PCR was performed with 1.25 U Taq DNApolymerase (Promega, Mannheim, Germany), 500 μMdNTPs and 26.7 pmol of the specific primers (OperonBiotechnologies, Cologne, Germany; Table 4). Annealingtemperature for all primers was 60°C. Amplification effi-ciencies for all primer pairs were determined by templatedilution series. Calculation of the relative mRNA concen-tration was made using the amplification efficiencies andthe Ct values [64]. Glyceraldehyde 3-phosphate dehydro-

36



genase (GAPDH) was used as housekeeping gene for nor-malization.

Enzyme assaysSOD in liver cytosol was determined with pyrogallol asthe substrate [65]. One unit of SOD activity is defined asthe amount of enzyme required to inhibit the autoxida-tion of pyrogallol by 50%. The activity of GSH-Px in livercytosol was determined with t-butyl hydroperoxide assubstrate [66]. One unit of GSH-Px activity is defined asone μmol reduced β-nicotinamide adenine dinucleotidephosphate oxidized per min. The activity of GST wasdetermined using 1-chloro-2,4-dinitrobenzene as sub-strate [67]. One unit of GST is defined as one nmol sub-strate consumed per min. Catalase activity in liverhomogenate was determined using hydrogen peroxide assubstrate [68]. One unit of catalase activity is defined asthe amount consuming 1 mmol hydrogen peroxide permin. GSH concentration in liver homogenates was deter-mined according to Griffith [69].

Determination of conjugated dienes, TBARS, α-tocopherol, cholesterol and triglycerides in the liverLipids from liver were extracted using a mixture of n-hex-ane and isopropanol (3:2, v/v). After drying the lipidextracts, 1 mg of extract was dissolved in 1 ml n-hexane.The concentrations of conjugated dienes were calculatedby using the molar extinction coefficient for conjugateddienes at 234 nm (ε = 29,500 mol × cm-1). The concentra-tions of TBARS were measured with thiobarbituric acid asreagent in a fluorimetric assay [70]. Concentration of α-tocopherol in liver tissue was determined by HPLC withfluorescence detection [70]. For determination of triglyc-erides and total cholesterol, an aliquot of the lipid extract

was dried, and the dried lipids were dissolved with TritonX-100 [71]. The concentrations of cholesterol and triglyc-erides were determined using enzymatic kits (Cat.-No.113009990314 for cholesterol and Cat.-No.157609990314 for triglycerides, Ecoline S+, DiaSys,Holzheim, Germany).

Determination of H2O2To determine the H2O2 content in liver homogenates, amethod [72] described for cell culture systems was modi-fied, using dihydrorhodamine 123 (DHR) as substrate.Homogenates were incubated with 27.5 μM DHR for 1 hat 37°C in a final volume of 400 μl. After incubation, thefluorescence of rhodamine 123, the oxidation product ofDHR, was measured (excitation wavelength: 485 nm,emission wavelength: 538 nm). As previously shown [73],this test is specific for H2O2 as DHR is specifially oxidizedby H2O2.

Transmission electron microscopySmall liver segments of three animals per group were fixedimmediately after dissection of the liver with 3% glutaral-dehyde in 0.1 M sodium cacodylate buffer (SCB, pH 7.2)for 3 hours at room temperature, washed with SCB, post-fixed with 1% osmiumtetroxide in SCB, dehydrated in agraded ethanol series, and embedded in epoxy resin [74].The material was sectioned with an ultramicrotome S(Leica, Bensheim, Germany). Ultrathin sections (80 nm)were transferred to coated copper grids and poststainedwith uranyl acetate and lead citrate. The sections wereobserved with an EM 900 transmission electron micro-scope (Zeiss SMT, Oberkochen, Germany) at an accelera-tion voltage of 80 kV. Electron micrographs were takenwith a slow scan camera (TRS, Dünzelbach, Germany).

Table 4: Sequences of specific primers used for RT-PCR

Gene (NCBI Genbank) Forward Primer Reverse Primer Size, bp

ACO (AF185048) CTCGCAGACCCAGATGAAAT TCCAAGCCTCGAAGATGAGT 218bax (AJ606301) CGAACTGATCAGGACCATCA ACAGCCCATCTTCTTCCAGA 190bcl-XL (NM_214285) GAAACCCCTAGTGCCATCAA GGGACGTCAGGTCACTGAAT 196Catalase (NM_214301) CAGCTTTAGTGCTCCCGAAC AGATGACCCGCAATGTTCTC 180c-fos (Y14808) CTGACACACTCCAAGCGGTA CTTCTCCTTCAGGTTGG 209c-jun (NM_213880) CAGAGCATGACCCTGAACCT TTCTTGGGGCATAGGAACTG 200c-myc (NM_001005154) AATGTCTTGGAACGCCAGAG CAACTGTTCTCGCCTCTTCC 204GAPDH (AF017079) AGGGGCTCTCCAGAACATCA

TCCTCGCGTGCTCTTGCTGGGGTTGG

446

GSH-Px (NM_214201) CAAGAATGGGGAGATCCTGA GATAAACTTGGGGTCGGTCA 190GST (NM_214300) TTTTTGCCAACCCAGAAGAC GGGGTGTCAAATACGCAATC 246p53 (NM_214145) GCGAGTATTTCACCCTCCAG TCAGGCCCTTCTCTCTTGAA 199PPARα (DQ437887) CAGCCTCCAGCCCCTCGTC GCGGTCTCGGCATCTTCTAG

G381

SOD (AF396674) TCCATGTCCATCAGTTTGGA CTGCCCAAGTCATCTGGTTT 250TNFα (X57321) CCCCTGTCCATCCCTTTATT AAGCCCCAGTTCCAATTCTT 200

Abbreviations: ACO, acyl-CoA oxidase; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; GSH-Px, glutathione peroxidase; GST, glutathione S-transferase; SOD, superoxide dismutase; TNFα, tumor necrosis factor α.

37

http://www.ncbi.nih.gov/entrez/query.fcgi?db=Nucleotide&cmd=search&term=AF185048

http://www.ncbi.nih.gov/entrez/query.fcgi?db=Nucleotide&cmd=search&term=AJ606301

http://www.ncbi.nih.gov/entrez/query.fcgi?db=Nucleotide&cmd=search&term=NM_214285


http://www.ncbi.nih.gov/entrez/query.fcgi?db=Nucleotide&cmd=search&term=Y14808







http://www.ncbi.nih.gov/entrez/query.fcgi?db=Nucleotide&cmd=search&term=DQ437887


http://www.ncbi.nih.gov/entrez/query.fcgi?db=Nucleotide&cmd=search&term=X57321



Peroxisomes were counted in 1,000 different regions perliver sample for each animal at a screen (12,000-fold mag-nification).

StatisticsThe results were analyzed using Minitab (State College,Pa, USA) statistical software (release 13). Statistical signif-icance of differences between control group and treatmentgroup was evaluated using Student's t-test. Mean valueswere considered significantly different for P < 0.05.

List of abbreviationsACO, acyl-CoA oxidase; DHR, dihydrorhodamine;GAPDH, glyceraldehyde 3-phosphate dehydrogenase;GSH-Px, glutathione peroxidase; GST, glutathione S-transferase; NF-KB, nuclear factor KB; PPARα, peroxisomeproliferator-activated receptor α; PPs, peroxisome prolif-erators; SCB, sodium cacodylate buffer; SOD, superoxidedismutase; TBARS, thiobarbituric acid-reactive sub-stances; TNFα, tumor necrosis factor α.

Authors' contributionsSL and BG carried out the feeding experiments, performedthe analyses and helped to draft the manuscript.

GH determined peroxisome count in the liver by trans-mission electrone microscopy.

HK participated in the design of the study and in the inter-pretation of the results and supervised the animal experi-ment.

KE conceived the study and its design, coordinated work,participated in the interpretation of the results, and pre-pared the manuscript.

All authors read and approved the final manuscript.

AcknowledgementsThe authors thank Bettina König for critical discussion of the manuscript. S. L. was supported by a grant from Land Sachsen-Anhalt.

References1. Schoonjans K, Staels B, Auwerx J: The peroxisome proliferator

activated receptors (PPARs) and their effects on lipidmetabolism and adipocyte differentiation. Biochim Biophys Acta1996, 1302:93-109.

2. Mandard S, Muller M, Kersten S: Peroxisome proliferator-acti-vated receptor α target genes. Cell Mol Life Sci 2004, 61:393-416.

3. Reddy JK, Rao S, Moody DE: Hepatocellular carcinomas in acat-alasemic mice treated with nafenopin, a hypolipidemic per-oxisome proliferator. Cancer Res 1976, 36:1211-1217.

4. Reddy JK, Azarnoff DL, Hignite CE: Hypolipidaemic hepatic per-oxisome proliferators form a novel class of chemical carcin-ogens. Nature 1980, 283:397-398.

5. Gonzalez FJ, Peters JM, Cattley RC: Mechanism of action of thenongenotoxic peroxisome proliferators: role of the peroxi-some proliferator-activator receptor-α. J Natl Cancer Inst 1998,90:1702-1709.

6. Peters JM, Cheung C, Gonzalez FJ: Peroxisome proliferator-acti-vated receptor-α and liver cancer: where do we stand? J MolMed 2005, 83:774-785.

7. Yeldandi AV, Rao MS, Reddy JK: Hydrogen peroxide generationin peroxisome proliferator-induced oncogenesis. Mutat Res2000, 448:159-177.

8. Ammerschlaeger M, Beigel J, Klein K-U, Müller SO: Characteriza-tion of the species-specifity of peroxisome proliferators inrats anf human hepatocytes. Toxicol Sci 2004, 78:229-240.

9. Nemali MR, Reddy MK, Usuda N, Reddy PG, Comeau LD, Rao MS,Reddy JK: Differential induction and regulation of peroxiso-mal enzymes: predictive value of peroxisome proliferation inidentifying certain nonmutagenic carcinogens. Toxicol ApplPharmacol 1989, 97:72-87.

10. Conway JG, Popp JA: Effect of the hepatocarcinogenic peroxi-some proliferator Wy-14,643 in vivo: no increase in ethaneexhalation or hepatic conjugated dienes. Toxicol Appl Pharmacol1995, 135:229-236.

11. Conway JG, Tomaszewski KE, Olson MJ, Cattley RC, Marsman DS,Popp JA: Relationship of oxidative damage to the hepatocar-cinogenicity of the peroxisome proliferators di(2-ethyl-hexyl)phthalate and Wy-14,643. Carcinogenesis 1989,10:513-519.

12. Kasai H, Okada Y, Nishimura S, Rao MS, Reddy JK: Formation of 8-hydroxydeoxyguanosine in liver DNA of rats following long-term exposure to a peroxisome proliferators. Cancer Res1989, 49:2603-2605.

13. Takagi A, Sai K, Umemura T, Hasegawa R, Kurokawa Y: Relation-ship between hepatic peroxisome proliferation and 8-hydroxydeoxyguanosine formation in liver DNA of rats fol-lowing long-term exposure to three peroxisome prolifera-tors; di(2-ethylhexyl) phthalate, aluminium clofibrate andsimfibrate. Cancer Lett 1990, 53:33-38.

14. Roberts RA: Non-genotoxic hepatocarcinogenesis: suppres-sion of apoptosis by peroxisome proliferators. Ann N Y Acad Sci1996, 804:588-611.

15. Peters JM, Cattley RC, Gonzalez FJ: Role of PPARα in the mech-anism of action of the nongenotoxic carcinogen and peroxi-some proliferator Wy-14,643. Carcinogenesis 1997,18:2029-2033.

16. Roberts RA, James NH, Hasmall SC, Holden PR, Lambe K, MacdonaldN, West D, Woodyatt NJ, Whitcome D: Apoptosis and prolifera-tion in nongenotoxic carcinogenesis: species differences androle of PPARα. Toxicol Lett 2000, 112:49-57.

17. Eacho PI, Foxworthy PS, Johnson WD, Hoover DM, White SL:Hepatic peroxisomal changes induced by a tetrazole-substi-tuted alkoxyacetophenone in rats and comparison withother species. Toxicol Appl Pharmacol 1986, 83:430-437.

18. Orton TC, Adam HK, Bentley M, Holloway B, Tucker MJ: Clobu-zarit: species differences in the morphological and biochem-ical response of the liver following chronic administration.Toxicol Appl Pharmacol 1984, 73:138-151.

19. Hanefeld M, Kemmer C, Kadner E: Relationship between mor-phological changes and lipid-lowering action of p-chlorphe-noxyisobutyric acid (CPIB) on hepatic mitochondria andperoxisomes in man. Atherosclerosis 1983, 46:239-246.

20. De La Iglesia FA, Lewis JE, Buchanan RA, Macus EL, McMahon G:Light and electron microscopy of liver in hyperlipoproteine-mic patients under long-term gemfibrozil treatment. Athero-sclerosis 1982, 43:19-37.

21. Blumcke S, Schwartzkopff W, Lobeck H, Edmondson NA, PrenticeDE, Blane GF: Influence of fenofibrate on cellular and subcellu-lar liver structure in hyperlipidemic patients. Atherosclerosis1983, 46:105-116.

22. Lalwani ND, Reddy MK, Ghosh S, Barnard SD, Molello JA, Reddy JK:Induction of fatty acid β-oxidation and peroxisome prolifera-tion in the liver of rhesus monkeys by DL-040, a new hypoli-pidemic agent. Biochem Pharmacol 1985, 34:3473-3482.

23. Reddy JK, Lalwani ND, Oureshi SA, Reddy MK, Moehle CM: Induc-tion of hepatic peroxisome proliferation in nonrodent spe-cies, including primates. Am J Pathol 1984, 114:171-183.

24. Hoivik DJ, Qualls CW Jr, Mirabile RC, Cariello NF, Kimbrough CL,Colton HM, Anderson SP, Santostefano MJ, Morgan RJ, Dahl RR,Brown AR, Zhao Z, Mudd PN Jr, Oliver WB Jr, Brown HHR, MillerRT: Fibrates induce hepatic peroxisome and mitochondrialproliferation without overt evidence of cellular proliferation

38






























































and oxidative stress in cynomolgus monkeys. Carcinogenesis2004, 25:1757-1769.

25. Cheon Y, Nara TY, Band MR, Beever JE, Wallig MA, Nakamura MT:Induction of overlapping genes by fasting and a peroxisomeproliferator in pigs: evidence of functional PPARα in nonpro-liferating species. Am J Physiol Regul Integr Comp Physiol 2005,288:R1525-1535.

26. Barkett M, Gilmore TD: Control of apoptosis by Rel/NF-kappaBtranscription factors. Onocogene 1995, 18:6910-6924.

27. DeMartin R, Schmid JA, Hoferwarbinek R: The NF-kappaB/Relfamily of transcription factors in oncogenic transformationand apoptosis. Mutat Res 1999, 437:231-243.

28. Tharappel JC, Nalca A, Owens AB, Ghabrial L, Konz EC, Glauert HP,Spear BT: Cell proliferation and apoptosis are altered in micedeficient in the NF-kappaB p50 subunit after treatment withthe peroxisome proliferator ciprofibrate. Toxicol Sci 2003,75:300-308.

29. Glauert HP, Eyigor A, Tharappel JC, Cooper S, Lee EY, Spear BT:Inhibition of hepatocarcinogenesis by the deletion of the p50subunit of NF-KB in mice administered the peroxisome pro-liferator Wy-14,643. Toxicol Sci 2006, 90:331-336.

30. Rolfe M, James NH, Roberts RA: Tumour necrosis factor α (TNFα) suppresses apoptosis and induces DNA synthesis inrodent hepatocytes: a mediator of the hepatocarcinogenic-ity of peroxisome proliferators? Carcinogenesis 1997,18:2277-2280.

31. Holden PR, Hasmall SC, James NH, West DR, Brindle RD, GonzalezFJ, Peters JM, Roberts RA: Tumour necrosis factor α (TNFα):role in suppression of apoptosis by the peroxisome prolifer-ator nafenopin. Cell Mol Biol 2000, 46:29-39.

32. Ledwith BJ, Johnson TE, Wagner LK, Pauley CJ, Manam S, GallowaySM, Nichols WW: Growth regulation by peroxisome prolifer-ators: opposing activities in early and late G1. Cancer Res 1996,56:3257-3264.

33. Ledwith BJ, Manam S, Troilo P, Joslyn DJ, Galloway SM, Nichols WW:Activation of immediate-early gene expression by peroxi-some proliferators in vitro. Mol Carcinog 1993, 8:20-27.

34. Miller RT, Glover SE, Stewart WS, Corton JC, Popp JA, Cattley RC:Effect on the expression of c-met, c-myc and PPAR-alpha inliver and liver tumors from rats chronically exposed to thehepatocarcinogenic peroxisome proliferator WY-14,643.Carcinogenesis 1996, 17:1337-1341.

35. Cherkaoui-Malki M, Meyer K, Cao WQ, Latruffe N, Yeldandi AV, RaoMS, Bradfield CA, Reddy JK: Identification of novel peroxisomeproliferator-activated receptor α (PPARα) target genes inmouse liver using cDNA microarray analysis. Gene Expr 2001,9:291-304.

36. Yu XX, Odle J, Drackley JK: Differential induction of peroxiso-mal β-oxidation enzymes by clofibric acid and aspirin in pig-let tissues. Am J Physiol Regul Integr Comp Physiol 2001,281:R1553-1561.

37. Peffer PL, Lin X, Odle J: Hepatic β-oxidation and carnitinepalmitoyltransferase I in neonatal pigs after dietary treat-ments of clofibric acid, isoproterenol, and medium-chaintriglycerides. Am J Physiol Regul Integr Comp Physiol 2005,288:R1518-R1524.

38. Anthony LE, Schmucker DL, Mooney JS, Jones AL: A quantitativeanalysis of fine structure and drug metabolism in livers ofclofibrate-treated young adult and retired breeder rats. JLipid Res 1978, 19:154-165.

39. He WS, Nara TY, Nakamura MT: Delayed induction of delta-6and delta-5 desaturases by a peroxisome proliferators. Bio-chem Biophys Res Commun 2002, 299:832-838.

40. Deisseroth A, Dounce AL: Catalase: Physical and chemicalproperties, mechanism of catalysis, and physiologic role.Physiol Rev 1970, 50:319-375.

41. Dhaunsi GS, Singh I, Orak JK, Singh AK: Antioxidant enzymes inciprofibrate-induced oxidative stress. Carcinogenesis 1994,15:1923-1930.

42. Marsman DS, Goldsworthy TL, Popp JA: Contrasting hepatocyticperoxisome proliferation, lipofuscin accumulation and cellturnover for the hepatocarcinogens Wy-14,643 and clofibricacid. Carcinogenesis 1992, 13:1011-1017.

43. Sies H: Glutathione and its role in cellular functions. Free RadicBiol Med 1999, 27:916-921.

44. O'Brien ML, Cunningham ML, Spear BT, Glauert HP: Effects of per-oxisome proliferators on glutathione and glutathionerelated enzymes in rats and hamsters. Toxicol Appl Pharmacol2001, 171:27-37.

45. Seo KW, Kim KB, Kim YJ, Choi JY, Lee KT, Choi KS: Comparisonof oxidative stress and change of xenobiotic metabolizingenzymes induced by phtalates in rats. Food Chem Toxicol 2004,42:107-114.

46. Cai Y, Appelkvist EL, DePierre JW: Hepatic oxidative stress andrelated defenses during treatment of mice with acetylsali-cylic acid and other peroxisome proliferators. J Biochem Toxicol1995, 10:87-94.

47. Arnaiz SL, Travacio M, Llesuy S, Boveris A: Hydrogen peroxidemetabolism during peroxisome proliferation by fenofibrate.Biochim Biophys Acta 1995, 1272:175-180.

48. O'Brien ML, Twaroski TP, Cunningham ML, Glauert HP, Spear BT:Effects of peroxisome proliferators on antioxidant enzymesand antioxidant vitamins in rats and hamsters. Toxicol Sci 2001,60:271-278.

49. Glauert HP, Srinivasan S, Tatum VL, Chen LC, Saxon DM, Lay LT,Borges T, Baker M, Chen LH, Robertson LW, Chow CK: Effects ofthe peroxisome proliferators ciprofibrate and perfluoro-decanoic acid on hepatic cellular antioxidants and lipid per-oxidation in rats. Biochem Pharmacol 1992, 43:1353-1359.

50. Lutz W, Fulda S, Jeremias I, Debatin KM, Schwab M: MycN and IFN-gamma cooperate in apoptosis of human neuroblastomacells. Oncogene 1998, 17:339-346.

51. Prendergast GC: Mechanisms of apoptosis by c-Myc. Oncogene1999, 18:2967-2987.

52. Oltvai ZN, Milliman CL, Korsmeyer SJ: Bcl-2 heterodimerizes invivo with a conserved homolog, Bax, that accelerates pro-grammed cell death. Cell 1993, 74:609-619.

53. Van Gijssel HE, Maassen CB, Mulder GJ, Meerman JH: p53 proteinexpression by hepatocarcinogens in the rat liver and itspotential role in mitoinhibition of normal hepatocytes as amechanism of hepatic tumour promotion. Carcinogenesis 1997,18:1027-1033.

54. Muller M, Strand S, Hug H, Heinemann EM, Walczak H, Hofmann WJ,Stremmel W, Krammer PH, Galle PR: Drug-induced apoptosis inhepatoma cells is mediated by the CD95 (APO-1/Fas) recep-tor/ligand system and involves activation of wild-type p53. JClin Invest 1997, 99:403-413.

55. Canuto RA, Muzio G, Maggiora M, Trombetta A, Martinasso G,Autelli R, Costelli P, Bonelli G, Baccino FM: Apoptosis induced byclofibrate in Yoshida AH-130 hepatoma cells: role of HMG-CoA reductase. J Lipid Res 2003, 44:56-64.

56. Canuto RA, Muzio G, Maggiora M, Autelli R, Barbiero G, Costelli P,Bonelli G, Baccino FM: Rapid and extensive lethal action of clof-ibrate on hepatoma cells in vitro. Cell Death Differ 1997,4:224-232.

57. Canuto RA, Muzio G, Bonelli G, Maggiora M, Autelli R, Barbiero G,Costelli P, Brossa O, Baccino FM: Peroxisome proliferatorsinduce apoptosis in hepatoma cells. Cancer Detect Prev 1998,22:357-366.

58. Passilly P, Jannin B, Hassell SJ, Latruffe N: Human HepG2 and ratFao hepatic-derived cell lines show different responses tociprofibrate, a peroxisome proliferator: analysis by flowcytometry. Exp Cell Res 1996, 223:436-442.

59. Goll V, Viollon-Abadie C, Nicod L, Richert L: Peroxisome prolifer-ators induce apoptosis and decrease DNA synthesis inhepatoma cell lines. Hum Exp Toxicol 2000, 19:193-202.

60. Xiao S, Anderson SP, Swanson C, Bahnemann R, Voss KA, Stauber AJ,Corton JC: Activation of peroxisome proliferator-activatedreceptor alpha enhances apoptosis in the mouse liver. ToxicolSci 2006, 92:368-377.

61. Martinasso G, Oraldi M, Trombetta A, Maggiora M, Bertetto O,Canuto RA, Muzio G: Involvement of PPARs in Cell Prolifera-tion and Apoptosis in Human Colon Cancer Specimens andin Normal and Cancer Cell Lines. PPAR Res 2007 in press. ArticleID93416

62. Tharappel JC, Cunningham ML, Spear BT, Glauert HP: Differentialactivation of hepatic NF-kappaB in rats and hamsters by theperoxisome proliferators Wy-14,643, gemfibrozil, and dib-utyl phthalate. Toxicol Sci 2001, 62:20-27.

63. National Research Council (Ed): Nutrient Requirement of Swine. TenthRevised Edition Washington DC: National Academie of Sciences; 1998.

39

































































































Publish with BioMed Central and every scientist can read your work free of charge

"BioMed Central will be the most significant development for disseminating the results of biomedical research in our lifetime."

Sir Paul Nurse, Cancer Research UK

Your research papers will be:

available free of charge to the entire biomedical community

peer reviewed and published immediately upon acceptance

cited in PubMed and archived on PubMed Central

yours — you keep the copyright

Submit your manuscript here:http://www.biomedcentral.com/info/publishing_adv.asp

BioMedcentral



64. Pfaffl MW: A new mathematical model for relative quantifica-tion in real-time RT-PCR. Nucleic Acids Res 2001, 29:e45.

65. Marklund S, Marklund G: Involvement of the superoxide anionradical in the autoxidation of pyrogallol and a convenientassay for superoxide dismutase. Eur J Biochem 1974, 47:469-474.

66. Paglia DE, Valentine WN: Studies on the quantitative and qual-itative characterization of erythrocyte glutathione peroxi-dase. J Lab Clin Med 1967, 70:158-169.

67. Habig WH, Pabst MJ, Jakoby WB: Glutathione S-transferases.The first enzymatic step in mercapturic acid formation. J BiolChem 1974, 249:7130-7139.

68. Aebi HE: Catalase. In Methods of Enzymatic Analysis, Band 3 3rd edi-tion. Edited by: Bergmeyer HU. Weinheim, Germany: VCH Verlags-gesellschaft mbH; 1986:273-286.

69. Griffith OW, Meister A: Origin and turnover of mitochondrialglutathione. Proc Natl Acad Sci USA 1985, 82:4668-4672.

70. Brandsch C, Ringseis R, Eder K: High dietary iron concentrationsenhance the formation of cholesterol oxidation products inthe liver of adult rats fed salmon oil with minimal effects onantioxidant status. J Nutr 2002, 132:2263-2269.

71. De Hoff JL, Davidson JH, Kritchevsky V: An enzymatic assay fordetermining free and total cholesterol in tissues. Clin Chem1978, 24:433-435.

72. Royall JA, Ischiropoulos H: Evaluation of 2',7'-dichlorofluorescinand dihydrorhodamine 123 as fluorescent probes for intrac-ellular H2O2 in cultured endothelial cells. Arch Biochem Biophys1993, 302:348-55.

73. Walrand S, Valeix S, Rodriguez C, Ligot P, Chassagne J, Vasson MP:Flow cytometry study of polymorphonuclear neutrophil oxi-dative burst: a comparison of three fluorescent probes. ClinChim Acta 2003, 331:103-110.

74. Spurr AR: A low-viscosity epoxy resin embedding medium forelectron microscopy. J Ultrastruct Res 1969, 26:31-43.

40

























http://www.biomedcentral.com/info/publishing_adv.asp


Feeding of a deep-fried fat causes PPARa activation in the liver of pigs

as a non-proliferating species

Sebastian Luci1, Bettina Konig1, Beatrice Giemsa1, Stefanie Huber1, Gerd Hause2, Holger Kluge1,

Gabriele I. Stangl1 and Klaus Eder1*1Institute of Agricultural and Nutritional Sciences, Martin-Luther-University of Halle-Wittenberg, Emil-Abderhalden-Strasse 26,

D-06108 Halle (Saale), Germany2Biocenter, Martin-Luther-University of Halle-Wittenberg, Weinbergweg 22, D-06120 Halle (Saale), Germany

(Received 24 August 2006 – Revised 20 November 2006 – Accepted 8 December 2006)

Recent studies have shown that dietary oxidised fats influence the lipid metabolism in rats by activation of PPARa. In this study, we investigated

whether a mildly oxidised fat causes activation of PPARa in pigs which are non-proliferators like man. Eighteen pigs were assigned to two groups

and received either a diet containing 90 g/kg of a fresh fat or the same diet with 90 g/kg of an oxidised fat prepared by heating for 24 h at 1808C in a

deep fryer. Pigs fed the oxidised fat had a higher peroxisome count, a higher activity of catalase and a higher mRNA concentration of mitochon-

drial 3-hydroxy-3-methylglutaryl-CoA synthase in the liver and a higher concentration of 3-hydroxybutyrate in plasma than pigs fed the fresh fat

(P,0·05). Hepatic mRNA concentrations of acyl-CoA oxidase and carnitine palmitoyltransferase-1 tended to be increased in pigs fed the oxidised

fat compared to pigs fed the fresh fat (P,0·10). Pigs fed the oxidised fat, moreover, had higher mRNA concentrations of sterol regulatory element-

binding protein (SREBP)-1 and its target genes acetyl-CoA carboxylase and stearoyl-CoA desaturase in the liver and higher mRNA concentrations

of SREBP-2 and its target genes 3-hydroxy-3-methylglutary-CoA reductase and LDL receptor in liver and small intestine. In conclusion, this study

shows that even a mildly oxidised fat causes activation of PPARa in the liver of pigs. Up-regulation of SREBP and its target genes in liver and

small intestine suggests that the oxidised fat could stimulate synthesis of cholesterol and TAG in these tissues.

Oxidised fat: Pig: PPARa: Cholesterol: Triacylglycerols

The typical western diet contains large quantities of PUFAthat are heated or processed to varying degrees. In fast-foodrestaurants fat is heated in fryers for up to 18 h daily, at tem-peratures close to 1808C (Frankel et al. 1984). Several studieswith animals have been performed to investigate the effects ofoxidised fats on the metabolism (reviewed in Cohn, 2002).Recently, it has been shown in rats that oxidised fats areable to influence the lipid metabolism by activation ofPPARa (Chao et al. 2001, 2004, 2005; Sulzle et al. 2004), atranscription factor belonging to the nuclear hormone receptorsuperfamily (Schoonjans et al. 1996). This is probably due tothe occurrence of hydroxy- and hydroperoxy fatty acids suchas hydroxy octadecadienoic acid and hydroperoxy octadeca-dienoic acid which are potent activators of PPARa (Deleriveet al. 2000; Mishra et al. 2004; Konig & Eder, 2006). Acti-vation of PPARa leads to an increase in the transcription ofgenes related to fatty acid transport across the cell membrane,intracellular lipid trafficking, mitochondrial and peroxisomalfatty acid uptake, and both mitochondrial and peroxisomalfatty acid b-oxidation, gluconeogenesis and ketogenesis

(Mandard et al. 2004). Recently, it has been shown thatPPARa activation influences also the expression or the proteo-lytic activation of sterol regulatory element-binding proteins(SREBP), transcription factors which control fatty acid syn-thesis and cholesterol homeostasis (Patel et al. 2001; Guoet al. 2001; Knight et al. 2005; Konig et al. 2006). Therefore,PPARa activation stimulates not only the degradation of fattyacids by enhancing b-oxidation but affects also the synthesisof cholesterol and TAG. Reduced liver and plasma concen-trations of TAG and cholesterol are typical effects observedin animals treated with PPARa agonists, and such effectshave been also observed in rats administered oxidised fats(Huang et al. 1988; Eder & Kirchgessner, 1998; Eder, 1999;Chao et al. 2001, 2004, 2005; Sulzle et al. 2004).

Regarding the expression of PPARa in tissues and theeffects of PPARa activation on transcription of its targetgenes, there are great differences between various species.In rodents, PPARa is highly expressed, and activation ofPPARa not only induces many genes involved in variousmetabolic pathways such as b-oxidation, ketogenesis and

*Corresponding author: Prof. Dr K. Eder, fax þ49 345 5527124, email [email protected]

Abbreviations: ACC, acetyl-CoA carboxylase; ACO, acyl-CoA oxidase; CPT-1, carnitine palmitoyltransferase-1; CYP7, cholesterol 7a-hydroxylase; HMG-CoA-R,

3-hydroxy-3-methylglutaryl-CoA reductase; L-FABP, liver fatty acid binding protein; mHMG-CoA-S, mitochondrial 3-hydroxy-3-methylglutaryl-CoA synthase;

MTP, microsomal TAG transfer protein; SCD, stearoyl-CoA desaturase; SOD, superoxide dismutase; SREBP, sterol regulatory element-binding protein; TBARS,

thiobarbituric acid-reactive substances.

British Journal of Nutrition (2007), 97, 872–882 doi: 10.1017/S0007114507669256q The Authors 2007

41

gluconeogenesis but also causes severe peroxisome prolifer-ation in the liver (Peters et al. 2005). In contrast to rodents,PPARa agonists do not induce peroxisome proliferation inthe liver of many other species, such as guinea pigs, swine,monkeys and man (Holden & Tugwood, 1999). These non-proliferating species have a lower expression of PPARa inthe liver and the response of many genes to PPARa activationis much weaker than in proliferating species. For that reason,effects related to PPARa activation observed in rodentscannot be directly applied for non-proliferating species suchas man. Therefore, it remains unknown whether oxidisedfats are able to cause PPARa activation also in non-proliferat-ing species.

The aim of the present study was to investigate whether adietary oxidised fat, prepared by heating sunflower oil underusual deep-frying conditions (1808C) for 24 h in a deepfryer, is able to activate PPARa and to cause peroxisome pro-liferation in pigs. Pigs have been chosen as a model since theybelong – like man – to the non-proliferating species (Yu et al.2001; Peffer et al. 2005) and since pig liver cells show a simi-larity to human liver cells in the gene response to PPARa ago-nists (Cheon et al. 2005). We focused our analyses on liverand small intestine as both tissues exhibit a high expressionof PPARa (Braissant et al. 1996; Lemberger et al. 1996).Moreover, both tissues play an important role in whole bodylipid homeostasis, i.e. synthesis and secretion of lipoproteinsrich in TAG and cholesterol (Lindsay & Wilson, 1965;Dietschy et al. 1993). We examined the expression of variousgenes involved in lipid metabolism which have been alreadyshown to be influenced by PPARa activation. Furthermore,in both tissues we determined gene expression of SREBPand important SREBP target genes involved in fatty acidsynthesis and cholesterol uptake and synthesis.

Materials and methods

Animals

For the experiment, eighteen male 8-week-old crossbred pigs((German Landrace £ Large White) £ Pietrain) were kept ina room under controlled temperature at 23 ^ 28C and55 ^ 5 % relative humidity with light from 06.00 to 18.00hours. One day before the beginning of the experimentalfeeding period, the pigs were weighed and randomly allocatedto two groups with body weights of 12·0 (SD 1·1) kg in thecontrol group and 12·2 (SD 0·9) kg in the treatment group.All experimental procedures described followed establishedguidelines for the care and use of laboratory animals andwere approved by the local veterinary office.

Diets and feeding

Both groups of pigs received a nutritionally adequate diet forgrowing pigs containing (in g/kg) wheat (400), soyabean meal(230), wheat bran (150), barley (100), sunflower oil or test oil(90), and mineral premix including L-lysine, DL-methionineand L-threonine (30). This diet contained 14·4 MJ metabolis-able energy and 185 g crude protein/kg. Diet intake was con-trolled, and each animal in the experiment was offeredan identical amount of diet per day. During the feedingperiod, the amount of diet offered each day was increased

continuously from 400 to 1200 g. The pigs had free accessto water via nipple drinking systems. The experimental dietswere administered for 28 d.

Preparation of the test fats

To prepare the oxidised fat, sunflower oil obtained from alocal supermarket was heated at a temperature of 1808C for24 h in a deep fryer. This treatment caused a loss of PUFAand tocopherols. The major fatty acids in the fresh and theoxidised fat, respectively, were (g/100 g total fatty acids): pal-mitic acid (16 : 0), 6·30 v. 6·70; stearic acid (18 : 0), 4·0 v. 4·2;oleic acid (18 : 1n-9), 22·8 v. 23·8; linoleic acid (18 : 2n-6),63·6 v. 59·9. Other fatty acids were present only in smallamounts (,0·5 g/100 g fatty acids). To equalise the fattyacid composition of the fresh and the oxidised fat, the freshfat was composed of a mixture of sunflower oil and palmoil (93 : 7, w/w). To adjust dietary vitamin E concentrations,we analysed the native concentrations of tocopherols in thefresh fat and in the oxidised fats after the thermal treatment.With consideration of the native tocopherol concentrationsof the dietary fats, the diets were supplemented indivi-dually with all-rac-a-tocopheryl acetate (the biopotency ofall-rac-a-tocopheryl acetate is considered to be 67 % of thatof a-tocopherol). The final vitamin E concentration was620 mg a-tocopherol equivalents/kg in both fats. Concen-trations of lipid peroxidation products were determined afterthe fats have been already included into the diets. Therefore,lipids of the diets were extracted by n-hexane and isopropanol(3 : 2, v/v; Hara & Radin, 1978). Concentration of thiobarbitu-ric acid-reactive substances (TBARS; Sidwell et al. 1954),conjugated dienes (Recknagel & Glende, 1984), peroxidevalue (Deutsche Gesellschaft fur Fettwissenschaft, 1994),acid value (Deutsche Gesellschaft fur Fettwissenschaft,1994) and concentration of total carbonyls (Endo et al.2001) were determined in the extracted fat.

Sample collection

After completion of the feeding period the animals were killedunder light anaesthesia. Each pig was fed its respective diet 4 hbefore being killed. After killing, blood was collected intoheparinised polyethylene tubes. Plasma was obtained bycentrifugation of the blood (1100 g, 10 min, 48C). Plasma lipo-proteins were separated by step-wise ultracentrifugation(Mikro-Ultrazentrifuge; Sorvall Products, Bad Homburg,Germany) at 900 000 g at 48C for 1·5 h. Plasma densitieswere adjusted by sodium chloride and potassium bromideand the lipoprotein fractions d , 1·006 kg/l VLDL plus chylo-microns, 1·006 , d , 1·063 kg/l LDL and d . 1·063 kg/lHDL were removed by suction. The liver was dissected andweighted and samples were stored at 2808C until analysis.For preparation of liver homogenate, 1 g liver tissue was hom-ogenised in PBS by TissueLyser (Qiagen, Haan, Germany),centrifuged at 600 g for 10 min at 48C and the supernatantwas stored at 2208C until analysis. For isolation of intestinalepithelial cells, the abdomen was immediately opened afterkilling, and a 35 cm intestinal segment was dissected startingat 30 cm distal to the pyloric sphincter, and flushed twicewith ice-cold wash buffer (PBS containing 0·2 mM-phenyl-methylsulphonyl fluoride and 0·5 mM-dithiothreitol, pH 7·4).

Effect of a deep-fried fat in pigs 873

42

The isolation of porcine intestinal epithelial cells was per-formed by the modified distended intestinal sac techniqueaccording to Fan et al. (2004). In brief, the intestinal segmentswere filled with 100 ml preincubation buffer (PBS containing27 mM-sodium citrate, 0·2 mM-phenylmethylsulphonyl fluorideand 0·5 mM-dithiothreitol, pH 7·4), sealed with strings and filledintestinal segments were incubated in a saline bath (154 mM-NaCl) for 15 min at 378C. Afterwards, the pre-incubationbuffer was discarded, and the intestinal segments were filledwith isolation buffer (PBS containing 1·5 mM-Na2EDTA,0·2 mM-phenylmethylsulphonyl fluoride, 0·5 mM-dithiothreitoland 2 mM-D-glucose, pH 7·4). Two major cell fractions, con-sisting of the upper and the crypt cell fraction, were sequen-tially isolated from intestinal segments through twoconsecutive incubations with isolation buffer at 378C for 40(upper cell fraction) and 60 min (crypt cell fraction), respect-ively. Each cell fraction was collected separately, and washedtwice with ice-cold PBS. Afterwards, cells were retained bycentrifugation (400 g, 4 min, 48C) and immediately frozen at2808C. For further analysis, we used the crypt cell fractionas it has been shown that these cells have a 6–8-fold highercapacity of lipid synthesis than villus cells (Shakir et al. 1978).

Lipid analysis

Lipids from liver were extracted with a mixture of n-hexaneand isopropanol (3 : 2, v/v; Hara & Radin, 1978). For determin-ation of the concentrations of lipids in liver, aliquots of the lipidextracts were dried and the lipids were dissolved using TritonX-100 (De Hoff et al. 1978). Concentrations of TAG and choles-terol in plasma and lipoproteins and those of liver were deter-mined using enzymatic reagent kits (cat. no. 113009990314for cholesterol and cat. no. 157609990314 for TAG; EcolineSþ, DiaSys, Holzheim, Germany).

Preparation of liver microsomal and cytosolic fractions

Liver (1 g) was homogenised in 10 ml 0·1 M-phosphate buffer,pH 7·4, containing 0·25 M-sucrose using a Potter-Elvehjemhomogeniser. Homogenates were centrifuged at 1000 g for10 min at 48C, and the supernatant was centrifuged at15 000 g for a further 15 min. The microsomal pellet wasobtained by centrifugation of the 15 000 g supernatant at105 000 g for 60 min. The resulting cytosolic fraction in thesupernatant was separated, microsomal pellets were resus-pended in the homogenisation buffer and all samples werestored at 2208C for further analysis. The protein concen-trations of cytosolic and microsomal fractions were deter-mined with the BCA reagent according to the protocol ofthe supplier (Interchim, Montelucon, France) using bovineserum albumin as standard.

RT–PCR analysis

Total RNA from liver tissue and enterocytes, respectively, wasisolated by the TissueLyser (Qiagen) using Trizol reagent(Invitrogen, Karlsruhe, Germany) according to the manufac-turer’s protocol. RNA concentration and purity were estimatedfrom the optical density at 260 and 280 nm (SpectraFluor Plus;Tecan, Crailsheim, Germany). The quality of all RNA sampleswas furthermore assessed by agarose gel electrophoresis.

Total RNA (1·2mg) was used for cDNA synthesis as describedpreviously (Konig & Eder, 2006). The mRNA concentrationof genes was measured by real-time detection PCR usingSYBRw Green I and the Rotor Gene 2000 system (CorbettResearch, Mortlake, Australia). Real-time detection PCRwas performed with 1·25 U Taq DNA polymerase, 500mM-dNTP and 26·7 pmol of the specific primers. For determinationof mRNA concentration a threshold cycle (Ct) and amplifica-tion efficiency was obtained from each amplification curveusing the software RotorGene 4·6 (Corbett Research). Calcu-lation of the relative mRNA concentration was made usingthe DDCt method as previously described (Pfaffl, 2001). Thehousekeeping gene glyceraldehyde-3-phosphate dehydrogen-ase was used for normalisation. The PCR primers used forreal-time RT–PCR were obtained from Operon (Koln,Germany) and Roth (Karlsruhe, Germany), respectively, andare listed in Table 1.

Enzyme assays

Superoxide dismutase (SOD) activity in liver cytosol wasdetermined according to the method of Marklund & Marklund(1974) with pyrogallol as the substrate. One unit of SODactivity is defined as the amount of enzyme required to inhibitthe autoxidation of pyrogallol by 50 %. The activity of gluta-thione peroxidase in liver cytosol was determined with t-butylhydroperoxide as substrate according to the method of Paglia& Valentine (1967). One unit of glutathione peroxidaseactivity is defined as 1mmol reduced b-nicotinamide adeninedinucleotide phosphate oxidised/min. The activity of gluta-thione S-transferase was determined using 1-chloro-2,4-dini-trobenzene as substrate as described by Habig et al. (1974).One unit of glutathione S-transferase is defined as one nmolsubstrate consumed/min. Catalase activity in liver homogenatewas determined using H2O2 as substrate according to themethod of Aebi (1986). One unit of catalase activity is definedas the amount consuming 1 mmol H2O2/min.

Determination of conjugated dienes, thiobarbituricacid-reactive substances and a-tocopherol

Lipids from liver were extracted using a mixture of n-hexaneand isopropanol (3 : 2, v/v; Hara & Radin, 1978). After dryingthe lipid extracts, 1 mg extract was dissolved in 1 ml n-hexane.The concentrations of conjugated dienes were calculated byusing the molar extinction coefficient for conjugated dienesat 234 nm (e ¼ 29 500 mol/cm). The concentrations ofTBARS were measured in liver homogenates as described(Brandsch et al. 2002). The concentration of a-tocopherol inliver tissue was determined by HPLC (Brandsch et al. 2002).

Determination of H2O2

To determine the H2O2 content in liver homogenates, themethod for cell culture systems described by Royall & Ischir-opoulos (1993) was modified, using dihydrorhodamine 123 assubstrate. Homogenates were incubated with 27·5mM-di-hydrorhodamine 123 for 1 h at 378C in a final volume of400ml. After incubation, the fluorescence of rhodamine 123,the oxidation product of dihydrorhodamine 123, was measured(excitation wavelength 485 nm, emission wavelength 538 nm).

S. Luci et al.874

43

Table

1.Characteristicsofthespecificprimers

usedforRT–PCR

analysis

Gene

Forw

ard

primer(from

50to

30 )

Reverseprimer(from

50to

30 )

bp

Annealingtemp.(8C)

NCBIGenBank

ACC

CTCCAGGACAGCACAGATCA

GCCGAAACATCTCTGGGATA

170

60

AF175308

ACO

CTCGCAGACCCAGATGAAAT

TCCAAGCCTCGAAGATGAGT

218

60

AF185048

apoCIII

GACACCTCCCTTCTGGACAA

TCCCAGAAGTCGGTGAACTT

185

60

NM_001002801

CPT-1

GCATTTGTCCCATCTTTCGT

GCACTGGTCCTTCTGGGATA

198

60

AF288789

CYP7

TATAGGGCACGATGCACAGA

ACCTGACCAGTTCCGAGATG

200

60

NM_001005352

FAS

GAACACGGCCTAGAAGTGGA

ATCTGGATCCTGCAGATGG

199

62

NM_213839

FATP

GGTTCCAGCCTGTTGAATGT

AACAAAACCTTGGTGCTTGG

275

60

DQ192231

FDPS

GAAAGGCAGGATTTCATCCA

AGAAGGCTTGGAGCAGTTCA

259

60

AY609787

GAPDH

AGGGGCTCTCCAGAACATCATCC

TCGCGTGCTCTTGCTGGGGTTGG

446

60

AF017079

Glutathioneperoxidase

CAAGAATGGGGAGATCCTGA

GATAAACTTGGGGTCGGTCA

190

60

NM_214201

GlutathioneS-transferase

TTTTTGCCAACCCAGAAGAC

GGGGTGTCAAATACGCAATC

246

60

NM_214300

HMG-C

oA-R

GGTCAGGATGCGGCACAGAACG

GCCCCACGGTCCCGATCTCTATG

127

65

S79678

I-FABP

TACAGCCTCGCAGACGGAACTG

TGCTTGATGAGGAGAGGAAAACAG

276

59

AY960624

Insig-1

AGAGGGAGTGGGCCAGTGTGATGC

ACGGGAGCCAGGAGCGGATGTAG

276

65

AY336601

Insig-2

AAATCACGCCAGCGCTAAAGTG

TCCTACTCCAAGGCCAAAACCAC

127

60

AY585269

LDLreceptor

AGAACTGGCGGCTGAAGAGCATC

GAGGGGTAGGTGTAGCCGTCCTG

115

60

AF118147

L-FABP

TTCGGTGCATGTCTAAGCTG

TGAGAGGGAGAGGATGAGGA

200

60

DQ182323

mAAT

TATGTCACCGTGCAGACCAT

CTCCTTCCACTGCTCAGGAC

309

60

M11732

mHMG-C

oA-S

GGACCAAACAGACCTGGAGA

ATGGTCTCAGTGCCCACTTC

198

62

U90884

MTP

CAGGACGGCAAAGAAAGAAGG

ATGGGAAGCAAAACCACAAGG

199

60

AY217034

NPC1

ACGCGGTATCTTTGGTCAAC

AGTGGCTCCCAGCAAGACTA

266

60

AF169635

NPC2

GGAGGGGAGGAGAAATCAAG

ATTCGGGTCTTGTCTGGTTG

267

60

NM_214206

PPARa

CAGCCTCCAGCCCCTCGTC

GCGGTCTCGGCATCTTCTAGG

382

58

DQ437887

SCD

ACGTTGTGCCAGTGAGTCAG

GTCTTGGCCTCTTGTGCTTC

206

62

NM_213781

SOD

TCCATGTCCATCAGTTTGGA

CTGCCCAAGTCATCTGGTTT

250

60

AF396674

SREBP-1

CCTCTGTCTCTCCTGCACC

ACAAAGAGAAGCGCCAAGAA

213

62

NM_214157

SREBP-2

CGCTCGCGAATCCTGCTGTG

GGTGCGGGTCCGTGTCGTG

103

65

DQ020476

ACC,acetyl-CoA

carboxylase;ACO,acyl-CoA

oxidase;CPT-1,carnitinepalm

itoyltransferase-1;CYP7,cholesterol7a-hydroxylase;FAS,fattyacid

synthase;FATP,fattyacid

transportprotein;FDPS,farnesyldiphosphate

synthase;

GAPDH,glyceraldehyde-3-phosphate

dehydrogenase;HMG-C

oA-R

,3-hydroxy-3-m

ethylglutaryl-CoA

reductase;I-FABP,intestinalfattyacid

binding

protein;Insig,insulin-induced

gene;L-FABP,liverfattyacid

binding

protein;

mAAT,mitochondrialaspartate

aminotransferase;mHMG-C

oA-S,mitochondrial3-hydroxy-3-m

ethylglutaryl-CoAsynthase;MTP,microsomalTAG

transferprotein;NPC,Niemann-PicktypeC;SCD,stearoyl-CoA

desaturase;SOD,

superoxidedismutase;SREBP,sterolregulatory

element-bindingprotein.


44

As previously shown by Walrand et al. (2003), dihydrorhoda-mine 123 is specifically oxidised by H2O2.

Determination of 3-hydroxybutyrate

Concentration of 3-hydroxybutyrate in plasma was determinedusing an enzymatic assay (cat. no. 10907979035; R-BiopharmAG, Darmstadt, Germany).

Transmission electron microscopy

Liver tissues were fixed in 3 % sodium cacodylate-bufferedglutaraldehyde (pH 7·2) and post-fixed with 1 % osmiumtetroxide. After washing three times, probes were dehydratedin an ethanol series and embedded in Spurr’s epoxy resin.For observations with an EM 900 transmission electron micro-scope (Carl Zeiss SMT, Oberkochen, Germany), ultrathinsections (80 nm) were mounted on copper grids. Catalase isknown to be located in peroxisomes specifically and wasmarked for a better visualisation of peroxisomes. For immuno-histochemistry, ultrathin sections were blocked for 30 minwith 1 % bovine serum albumin and 0·1 % Tween in PBSand incubated overnight with sheep polyclonal anti-catalaseserum (1 : 50; Biotrend, Koln, Germany). For detection of pri-mary antibody, sections were incubated for 1 h with a gold-marked donkey–anti-sheep antibody (1 : 25; Biotrend) andfinally stained with uranyl acetate/lead citrate. Peroxisomeswere counted in 1000 different prints per liver sample foreach animal with a magnification of 12 000£ .

Statistics

The results were analysed using Minitab (State College, PA,USA) statistical software (release 13). Statistical significanceof differences of the mean values of the two groups of pigswas evaluated using Student’s t test. Mean values were con-sidered significantly different for P,0·05.

Results

Fatty acid composition and concentration of lipidperoxidation products in the dietary fats

Palmitic, stearic, oleic and linoleic acid were the major fattyacids in the dietary fats. The sum of these fatty acidsaccounted for about 95 g/100 g total fatty acids in the fats(Table 2). Amounts of stearic, oleic and linoleic acid werenearly identical in both fats; the amount of stearic acid wasslightly higher in the fresh fat than in the oxidised fat. Per-oxide value, acid value and concentration of conjugateddienes were 4–5-fold higher in the oxidised than in thefresh fat included in the diet (Table 2). The concentration oftotal carbonyls was 10-fold higher and that of TBARS was30-fold higher in the oxidised than in the fresh fat (Table 2).

Body weights, antioxidant status and concentrations of lipidperoxidation products in the liver

Body weights of the pigs at the end of the experiment on day28 did not differ between the two groups (25·6 (SD 1·4) v. 26·0(SD 1·5) kg in pigs fed the oxidised fat v. pigs fed the fresh fat;

nine pigs per group). Pigs fed the oxidised fat had a highermRNA concentration and a higher activity of SOD and alower activity of microsomal glutathione S-transferase in theliver than pigs fed the fresh fat (P,0·05; Table 3). Activitiesof glutathione peroxidase and cytosolic glutathione S-transfer-ase as well as mRNA concentrations of these enzymes in theliver did not differ between both groups of pigs (Table 3).Concentrations of total, reduced and oxidised glutathione inthe liver also did not differ between the two groups of pigswhereas the concentration of a-tocopherol was lower in pigsfed the oxidised fat than in pigs fed the fresh fat (P,0·05;Table 3). Concentration of TBARS in the liver did not differbetween the two groups of pigs whereas the concentration ofconjugated dienes was slightly but significantly higher inpigs fed the oxidised fat than in pigs fed the fresh fat(P,0·05; Table 3).

Indices of peroxisome proliferation

Liver weights of the pigs were not different between the twogroups but pigs fed the oxidised fat had a higher peroxisomecount and a higher activity of catalase in the liver than pigsfed the fresh fat (P,0·05; Table 4). Relative mRNA concen-tration of acyl-CoA oxidase (ACO), a peroxisomal enzyme, inthe liver, was 34 % higher in pigs fed the oxidised fat than incontrol animals (P¼0·062; Table 4). The concentration ofH2O2 which is mainly released from peroxisomal oxidaseswas not different between the two groups of pigs (Table 4).

mRNA concentrations of genes in liver and intestine

In liver, mRNA concentrations of PPARa and genes involvedin fatty acid transport and oxidation [liver fatty acid bindingprotein (L-FABP), carnitine palmitoyltransferase-1 (CPT-1)],fatty acid and cholesterol synthesis [SREBP-1 and -2, insu-lin-induced gene-1 and -2, fatty acid synthase, acetyl-CoAcarboxylase (ACC), stearoyl-CoA desaturase (SCD), 3-hydroxy-3-methylglutaryl-CoA reductase (HMG-CoA-R)], cholesteroluptake (LDL receptor), bile acid synthesis [cholesterol 7a-hydroxylase (CYP7)], lipoprotein assembly and secretion[microsomal TAG transfer protein (MTP)], inhibition oflipoprotein lipase (apo CIII) and ketogenesis [mitochondrial3-hydroxy-3-methylglutaryl-CoA synthase (mHMG-CoA-S)]

Table 2. Major fatty acids and concentrations of some lipidperoxidation products in the fresh and the oxidised fat afterinclusion into the diet

Fresh fat Oxidised fat

Major fatty acids (g/100 g fatty acids)16 : 0 9·0 6·718 : 0 4·1 4·218 : 1n-9 23·7 23·818 : 2n-6 59·8 59·9

Peroxidation productsConjugated dienes (mmol/kg) 22·7 89·1TBARS (mmol/kg) 9 271Peroxide value (mEq O2/kg) 2·5 10·0Acid value (g KOH/kg) 1·6 8·0Total carbonyls (mmol/kg) 2·5 24·5

TBARS, thiobarbituric acid-reactive substances.

S. Luci et al.876

45

were determined (Fig. 1). Pigs fed the oxidised fat had signifi-cantly higher mRNA concentrations of mHMG-CoA-S, a clas-sical PPARa target gene, SREBP-1 and its target genes ACCand SCD, and SREBP-2 and its target genes HMG-CoA-R andLDL receptor than control pigs fed the fresh fat (P,0·05).mRNA concentration of CYP7 was lower in pigs fed theoxidised fat than in pigs fed the fresh fat (P,0·05). mRNAconcentrations of CPT-1 and MTP, two other PPARa targetgenes, tended to be increased in pigs fed the oxidised fat(P¼0·074 and P¼0·065, respectively) compared to pigsfed the fresh fat whereas mRNA concentrations of PPARa,L-FABP, apo CIII, insulin-induced gene-1 and -2, and fattyacid synthase were not different between pigs fed the freshfat and those fed the oxidised fat (Fig. 1).

In enterocytes, relative mRNA concentrations of PPARaand of proteins involved in fatty acid uptake (L-FABP,intestinal fatty acid binding protein, fatty acid transportprotein, mitochondrial aspartate aminotransferase), fatty acidoxidation (ACO, CPT-1), intracellular trafficking of choles-terol (Niemann-Pick type C1 and 2) and fatty acid synthesis(SREBP-1, fatty acid synthase) were not different betweenpigs fed the oxidised fat and those fed the fresh fat (Fig. 2).

However, mRNA concentration of SREBP-2 and its targetgenes HMG-CoA-R and LDL receptor, involved in cholesterolsynthesis and uptake, were higher in pigs fed the oxidised fatthan in pigs fed the fresh fat (P,0·05; Fig. 2). mRNA concen-tration of farnesyl diphosphate synthase did not differ betweenthe two groups of pigs (Fig. 2).

Concentrations of TAG and cholesterol in liver, plasmaand lipoproteins

Concentrations of TAG in liver, plasma and TAG-rich lipo-proteins did not differ between pigs fed the fresh fat andthose fed the oxidised fat. Concentrations in pigs fed the oxi-dised fat v. pigs fed the fresh were (nine pigs per group): liver,88 (SD 20) v. 91 (SD 19) mmol/g; plasma, 0·96 (SD 0·26) v.1·09 (SD 0·17) mmol/l; chylomicrons þ VLDL, 0·80 (SD

0·25) v. 0·93 (SD 0·16) mmol/l. Concentrations of cholesterolin liver, plasma, LDL and HDL were also not differentbetween the two groups of pigs. Concentrations in pigsfed the oxidised fat v. pigs fed the fresh were: liver,73 (SD 14) v. 69 (SD 10) mmol/g; plasma, 2·63 (SD 0·32)

Table 3. mRNA concentrations and activities of antioxidant enzymes and concentrationsof antioxidants and lipid peroxidation products in livers of pigs fed a diet with a fresh fat oran oxidised fat

(Mean values and standard deviations)

Fresh fat (n 9) Oxidised fat (n 9)

Mean SD Mean SD

Superoxide dismutasemRNA concentration (relative) 1·00 0·22 1·24* 0·14Activity (U/mg protein) 42·7 8·4 58·8* 6·0

Glutathione S-transferase 202 42 144* 13mRNA concentration (relative) 1·00 0·31 1·27 0·23Activity in microsomes (U/mg protein) 202 42 144* 13Activity in cytosol (U/mg protein) 760 262 761 186

Glutathione peroxidasemRNA concentration (relative) 1·00 0·13 1·14 0·22Activity (U/mg protein) 4·72 0·77 5·14 0·75

Glutathione, total (nmol/mg) 2·13 0·51 2·17 0·37Glutathione, reduced (nmol/mg) 1·70 0·50 1·84 0·55Glutathione, oxidised (nmol/mg) 0·21 0·12 0·17 0·10a-Tocopherol (nmol/g) 14·5 2·5 11·9* 1·8Conjugated dienes (mmol/mg protein) 16 1 18* 3TBARS (mmol/g) 7·2 1·6 7·3 2·8

TBARS, thiobarbituric acid-reactive substances.Mean values were significantly different from those of the fresh fat group: *P,0·05.

Table 4. Indices of peroxisome proliferation in livers of pigs fed a diet with a fresh fat or an oxidised fat

(Mean values and standard deviations)

Fresh fat (n 9) Oxidised fat (n 9)

Mean SD Mean SD

Liver weight (g) 673 63 700 64Peroxisome count (number/print) 366 67 515* 91Acyl-CoA oxidase mRNA concentration (relative) 1·00 0·33 1·34† 0·37Catalase (U/mg protein) 0·75 0·14 0·89* 0·13H2O2 (fluorescence/g liver) 29 372 12 343 29 437 8361

Mean values were significantly different from those of the fresh fat group: †P,0·1; *P,0·05.


46

v. 2·83 (SD 0·22) mmol/l; LDL, 0·96 (SD 0·16) v. 0·97 (SD 0·15)mmol/l; HDL, 1·02 (SD 0·18) v. 1·13 (SD 0·11) mmol/l.

Concentration of 3-hydroxybutyrate in plasma

Pigs fed the oxidised fat had a higher concentration of3-hydroxybutyrate in plasma than pigs fed the fresh fat(1·23 (SD 0·58) v. 0·52 (SD 0·27) mmol/l; P,0·05).

Discussion

In the present study, pigs were fed a diet containing an oxi-dised fat prepared under usual deep-frying conditions. Therelatively low concentrations of lipid peroxidation products(conjugated dienes, TBARS, peroxides and carbonyls) in theoxidised fat indicate that this fat was mildly oxidised. Concen-trations of peroxidation products in this fat were indeed even

lower than in soyabean oil or hydrogenated animal–vegetableoil blends used for frying of potatoes at 1908C over a period of24 h (Frankel, 1998). The reason for the relatively low degreeof oxidation is that we did not add foodstuffs to be fried duringthe preparation of the oil as we wanted to avoid contaminationof the oil with food ingredients. It is well known that ingredi-ents of foodstuffs, i.e. metal ions, enhance the lipid peroxi-dation process during frying of fats (Kubow, 1992). Theconcentration of conjugated dienes which include the potentPPARa activators hydroxy- and hydroperoxy fatty acids(Delerive et al. 2000; Konig & Eder, 2006) was approximatelyfour times higher in the oxidised fat than in the fresh fat. Thefinding of an increased activity of SOD and a slightly elevatedconcentration of conjugated dienes, together with the obser-vation of a slightly reduced concentration of a-tocopherol,indicates that the oxidised fat produced oxidative stress inthe liver of the pigs. It has been demonstrated that under

** *

0

0.5

1

1.5

2

2.5

PPARαACO

CPT-1

L-FA

BP

I-FABP

FATP

mAAT

NPC-1

NPC-2

SREBP-1

SREBP-2

Insig

-1

Insig

-2FA

S

HMG-C

oA-RFD

PS

LDL r

ecep

tor

Fold

ove

r co

ntr

ol

Fig. 2. Relative mRNA concentrations (--- represents 1·00) of various genes involved in intestinal lipid metabolism in livers of pigs fed a diet with a fresh fat or an

oxidised fat. Values were determined by real-time detection RT–PCR using the mRNA concentration of glyceraldehyde-3-phosphate dehydrogenase for normali-

sation. Values are means with their standard deviations depicted by vertical bars (n 9) obtained for the pigs fed the oxidised fat relative to the values of the control

group fed fresh fat. ACO, acyl-CoA oxidase; CPT-1, carnitine palmitoyltransferase-1; FAS, fatty acid synthase; FATP, fatty acid transport protein; FDPS, farnesyl

diphosphate synthase; HMG-CoA-R, 3-hydroxy-3-methylglutaryl-CoA reductase; I-FABP, intestinal fatty acid binding protein; Insig, insulin-induced gene; L-FABP,

liver fatty acid binding protein; mAAT, mitochondrial aspartate aminotransferase; NPC, Niemann-Pick type C; SREBP, sterol regulatory element-binding protein.

Mean values were significantly different from those of the fresh fat group: *P , 0·05.

*

**

*

**

*

*

0

0.5

1

1.5

2

2.5

3

PPARαCPT-1

L-FA

BP

mHM

G-CoA-S

apo-C

III

SREBP-1

SREBP-2

Insig

-1

Insig

-2FA

SACC

SCD

HMG-C

oA-R

LDL r

ecep

tor

CYP7M

TP

Fold

ove

r co

ntr

ol

Fig. 1. Relative mRNA concentrations (--- represents 1·00) of various genes involved in hepatic lipid metabolism in enterocytes of pigs fed a diet with a fresh fat

or an oxidised fat. Values were determined by real-time detection RT–PCR using the mRNA concentration of glyceraldehyde-3-phosphate dehydrogenase for

normalisation Values are means with their standard deviations depicted by vertical bars (n 9) obtained for the pigs fed the oxidised fat relative to the values of the

control group fed fresh fat. ACC, acetyl-CoA carboxylase; CPT-1, carnitine palmitoyltransferase-1; CYP7, cholesterol 7a-hydroxylase; FAS, fatty acid synthase;

HMG-CoA-R, 3-hydroxy-3-methylglutaryl-CoA reductase; Insig, insulin-induced gene; L-FABP, liver fatty acid binding protein; mHMG-CoA-S, mitochondrial

3-hydroxy-3-methylglutaryl-CoA synthase; MTP, microsomal TAG transfer protein; SCD, stearoyl-CoA desaturase; SREBP, sterol regulatory element-binding

protein. Mean values were significantly different from those of the fresh fat group: *P , 0·05.

S. Luci et al.878

47

oxidative stress, expression of SOD is stimulated and concen-tration of a-tocopherol is reduced due to an enhancedconsumption (Liu & Huang, 1995; Ruiz-Gutierrez et al.1999; Atalay et al. 2000). However, the oxidative stressproduced by the oxidised fat was very moderate as concen-trations of TBARS and glutathione remained completelyunchanged. In rodents treated with PPARa agonists such asfibrates or WY-14,643, production of H2O2 is largelyincreased due to a strong up-regulation of peroxisomal oxi-dases, and this causes oxidative stress and contributes to hepa-tocarcinogenesis in these species (Peters et al. 2005). In thepresent study, feeding the oxidised fat did not increase theconcentration of H2O2 in the liver. This was probably due totwo reasons: first, there was only a slight increase in themRNA concentration of ACO, one of the enzymes producingH2O2; second, activity of catalase, the key enzyme ofdecomposition of H2O2 in peroxisomes was increased. There-fore, generation of H2O2 did not contribute to oxidative stressin animals treated with oxidised fat. The reason for the mod-erate oxidative stress may be that a part of the dietary lipidperoxidation products is absorbed in the intestine and reachesthe liver via lipoproteins (Staprans et al. 2005). Production ofoxidative stress by intake of strongly oxidised fats has beenshown several times in rodents (Yoshida & Kajimoto, 1989;Liu & Huang, 1996; Liu & Lee, 1998; Ammouche et al.2002; Eder et al. 2004; Keller et al. 2004a, b). The presentstudy shows for the first time that even a mildly oxidisedfat, as used in human nutrition, can induce moderate oxidativestress in pigs as a non-proliferating species.

To find out whether the mildly oxidised fat caused activationof PPARa in the liver of pigs, we determined mRNA concen-trations of the classical PPARa target genes ACO, CPT-1 andmHMG-CoA-S as well as peroxisome count, activity of catalaseand plasma concentration of 3-hydroxybutyrate. Recent studiesin pigs have shown that activation of PPARa in pigs, by eithertreatment with clofibrate or by fasting, leads to an increasedexpression of these PPARa target genes, and in turn stimulatesmitochondrial and peroxisomal b-oxidation and ketogenesis(Yu et al. 2001; Peffer et al. 2005; Cheon et al. 2005). The find-ing of an increased peroxisome count together with increasedactivity of catalase, a peroxisomal enzyme, a significantlyincreased mRNA concentration of mHMG-CoA-S and anincreased plasma concentration of 3-hydroxybutyrate stronglyindicate that the oxidised fat caused PPARa activation in theliver of the pigs. The finding that mRNA concentrations ofACO and CPT-1, two other classical PPARa target geneswere also increased by 34 and 29 %, although not significantlydifferent to control, supports the assumption that the oxidisedfat induced hepatic PPARa activation in the pigs. It has beenshown that these two enzymes are only moderately up-regulatedin pig liver by PPARa agonists. For instance, in pigs treated withclofibrate, a strong PPARa agonist, hepatic gene expression ofCPT-1 and ACO was only 1·89- and 1·42-fold, respectively, in-creased over control while gene expression of mHMG-CoA-Swas increased 3·32-fold (Cheon et al. 2005). This presents anexplanation for the observations that mHMG-CoA-S wassignificantly increased in pigs treated with oxidised fat andthat ACO and CPT-1 were only slightly increased. The findingthat mRNA concentration of MTP, a gene recently shown tobe up-regulated by PPARa activation (Ameen et al. 2005),tended to be increased in the liver of pigs fed the oxidised fat

also indicates that the oxidised fat caused PPARa activation inthe liver. Recently, studies in rats have already shown that oxi-dised fats are able to activate PPARa in the liver (Huang et al.1988; Chao et al. 2001; Sulzle et al. 2004). In these rat studies,up-regulation of PPARa target genes in the liver was muchstronger than in pigs of the present study. This may have twodifferent reasons: first, most PPARa target genes respond stron-ger to PPARa activation in rats than in non-proliferating speciessuch as pigs or man; second, fats used in the rat studies were morestrongly oxidised than the mildly oxidised fat used in the presentstudy. The present study shows for the first time that even a mildlyoxidised fat causes activation of PPARa in pigs which are, asman, less sensitive to PPARa agonists than rodents.

To study whether the oxidised fat caused PPARa activationin small intestine, we considered in addition to the classicalPPARa target genes ACO and CPT-1, several genes involvedin fatty acid transport (L-FABP, intestinal fatty acid bindingprotein, fatty acid transport protein and mitochondrial aspar-tate aminotransferase) and cholesterol trafficking (Niemann-Pick type C1 and 2) in intestinal tissue. All these geneshave been shown to be up-regulated by PPARa activation(Darimont et al. 1998; Motojima et al. 1998; Mochizukiet al. 2001; Chinetti-Gbaguidi et al. 2005). The finding thatnone of these genes was up-regulated in cells of small intes-tine indicates that oxidised fat caused no or even weakPPARa activation and does not influence intestinal fattyacid transport and cholesterol trafficking.

Synthesis of lipids in mammalian cells is controlled by a net-work involving the action of insulin-induced genes and SREBP,and it has been recently shown in several studies that this networkis influenced by PPARa activation (Guo et al. 2001; Patel et al.2001; Knight et al. 2005; Konig et al. 2006). The present studyshows that feeding a mildly oxidised fat increased the mRNA con-centration of SREBP-1 and its target genes ACC and SCD, twokey enzymes of de novo fatty acid synthesis, in the liver. Thesealterations may be caused by activation of PPARa in the liver.Knight et al. (2005) found that treatment with WY 14,643, a syn-thetic PPARa agonist, causes a strong up-regulation of enzymesinvolved in hepatic fatty acid synthesis and stimulates fatty acidsynthesis in wild-type mice but not in PPARa null mice. Knightet al. (2005) suggest that up-regulation of hepatic fatty acid syn-thesis is a compensatory response on the increased fatty acid oxi-dation to maintain a constant cellular TAG level. The finding thatTAG levels in liver and plasma were not reduced in pigs fed theoxidised fat compared to control pigs indeed suggests that anincreased b-oxidation of fatty acids was compensated by anincreased fatty acid synthesis. As there is no evidence for adirect action of PPARa on the promoter regions of SREBP-1and ACC genes, it is likely that the increased mRNA concen-trations of these genes are an indirect result of PPARa activation.In contrast, SCD is not only dependent on SREBP-1 but has also aPPAR response element in its promoter (Miller & Ntambi, 1996).Therefore, its transcription may have been in part directly stimu-lated by PPARa activation. An up-regulation of SCD which cat-alyses the formation of MUFA from SFA has also been observedin pigs treated with clofibrate (Cheon et al. 2005). These findingsof the effects of the oxidised fat on gene expression of lipogenicenzymes observed in pigs are opposite to those observed in ratsin which a dietary oxidised fat causes a down-regulation of lipo-genic enzymes and a strong reduction of liver and plasma TAG(Eder & Kirchgessner, 1998; Eder et al. 2003).


48

It is moreover shown that feeding the mildly oxidised fatled to a moderate but significant up-regulation of SREBP-2,and its target genes HMG-CoA-R and LDL receptor, in bothliver and small intestine. The present findings suggest thatthe oxidised fat could have stimulated synthesis and uptakeof cholesterol in these tissues. As this effect occurs not onlyin the liver but also in the small intestine where no PPARaactivation was found in pigs fed the oxidised fat, it is question-able whether these effects are linked to PPARa activation.The finding that hepatic genes involved in cholesterol syn-thesis were not altered in pigs treated with clofibrate indeedsuggests that PPARa activation does not influence SREBP-2controlled transcription of genes involved in cholesterolhomeostasis (Cheon et al. 2005). On the other hand, treatmentwith the PPARa agonist WY 14,643 caused an up-regulationof genes involved in hepatic cholesterol synthesis in wild-typemice but not in PPARa null mice, indicating that PPARa acti-vation indeed could directly stimulate cholesterol synthesis(Knight et al. 2005). It should be noted, however, that thereis also another study that found a suppression of geneexpression and proteolytic activation of SREBP-2, and astrong down-regulation of its target genes accompanied byreduced cholesterol synthesis in rats (Konig et al. 2006).The effect of PPARa activation on SREBP-2-dependentcholesterol synthesis is not yet clear and may also be differentbetween various species. Besides an up-regulation of genesinvolved in synthesis and uptake of cholesterol, we found adown-regulation of CYP7, the key enzyme of bile acid for-mation, in the liver. It has been shown in human and ratliver cells that PPARa agonists lower CYP7 expression prob-ably by reducing the availability of hepatic nuclear factor 4awhich is required for binding to a DR-1 in CYP7 promoter(Marrapodi & Chiang, 2000; Patel et al. 2000). Therefore,we assume that down-regulation of CYP7 in the liver ofpigs fed the oxidised fat was caused by PPARa activationinduced by the oxidised fat. Increased hepatic cholesterol syn-thesis and uptake of cholesterol into the liver, together with adecreased bile acid synthesis, is expected to increase hepaticcholesterol concentration. In contradiction to this, liver andplasma cholesterol concentrations were unchanged in pigsfed the oxidised fat compared to pigs fed the fresh fat. Weassume that the changes in gene expression were too small toinduce phenotypical alterations of cholesterol concentrations.

In conclusion, the present study shows that a mildly oxidisedfat causes PPARa activation in the liver of pigs as indicated byan increased peroxisome count, a moderate up-regulation ofPPARa target genes and a stimulation of ketogenesis. Moreover,the oxidised fat led to an up-regulation of the expressionof SREBP-1 and SREBP-2 and their target genes involved inTAG and cholesterol synthesis, suggesting a stimulation oflipid synthesis. As the fat used in the present study was evenless oxidised than fats used for deep-frying of foods, and asthere exists a similarity in the gene response to PPARa agonistsbetween pig and human liver cells, deep-fried fats could exertsimilar effects in man.

Acknowledgements

Sebastian Luci and Bettina Konig contributed equally to thiswork.

References

Aebi HE (1986) Catalase. In Methods of Enzymatic Analysis, 3rd ed.

pp. 273–286 [HU Bergemeyer, editor]. Weinheim, Germany: VCH

Verlagsgesellschaft mbH.

Ameen C, Edvardsson U, Ljungberg A, Asp L, Akerblad P, Tuneld A,

Olofsson SO, Linden D & Oscarsson J (2005) Activation of peroxi-

some proliferator-activated receptor alpha increases the expression

and activity of microsomal triglyceride transfer protein in the liver.

J Biol Chem 280, 1224–1229.

Ammouche A, Rouaki F, Bitam A & Bellal MM (2002) Effect of

ingestion of thermally oxidized sunflower oil on the fatty acid com-

position and antioxidant enzymes of rat liver and brain in develop-

ment. Ann Nutr Metab 46, 268–275.

Atalay M, Laaksonen DE, Khanna S, Kaliste KE, Hanninen O & Sen

CK (2000) Vitamin E regulates changes in tissue antioxidants

induced by fish oil and exercise. Med Sci Sports Exerc 32,

601–607.

Brandsch C, Ringseis R & Eder K (2002) High dietary iron concen-

trations enhance the formation of cholesterol oxidation products

in the liver of adult rats fed salmon oil with minimal effects on

antioxidant status. J Nutr 132, 2263–2269.

Braissant O, Foufelle F, Scotto C, Dauca M & Wahli W (1996)

Differential expression of peroxisome proliferator-activated re-

ceptors (PPARs): tissue distribution of PPAR-alpha, -beta, and

-gamma in the adult rat. Endocrinology 137, 354–366.

Chao P-M, Chao C-Y, Lin F-J & Huang C-J (2001) Oxidized frying

oil up-regulates hepatic acyl-CoA oxdase and cytochrome P450

4A1 genes in rats and activates PPARa. J Nutr 131, 3166–3174.

Chao PM, Hsu SC, Lin FJ, Li YJ & Huang CJ (2004) The up-regu-

lation of hepatic acyl-CoA oxidase and cytochrome P450 4A1

mRNA expression by dietary oxidized frying oil is comparable

between male and female rats. Lipids 39, 233–238.

Chao PM, Yang MF, Tseng YN, Chang KM, Lu KS & Huang CJ

(2005) Peroxisome proliferation in liver of rats fed oxidized

frying oil. J Nutr Vitaminol (Tokyo) 51, 361–368.

Cheon Y, Nara TY, Band MR, Beever JE, Wallig MA & Nakamura

MT (2005) Induction of overlapping genes by fasting and a peroxi-

some proliferator in pigs: evidence of functional PPARa in non-

proliferating species. Am J Physiol Regul Integr Comp Physiol

288, R1525–R1535.

Chinetti-Gbaguidi G, Rigamonti E, Helin L, Mutka AL, Lepore M,

Fruchart JC, Clavey V, Ikonen E, Lestavel S & Staels B (2005)

Peroxisome proliferator-activated receptor alpha controls

cellular cholesterol trafficking in macrophages. J Lipid Res 46,

2717–2725.

Cohn JS (2002) Oxidized fat in the diet, postprandial lipaemia and

cardiovascular disease. Curr Opin Lipidol 13, 19–24.

Darimont C, Gradoux N, Cumin F, Baum HP & De Pover A (1998)

Differential regulation of intestinal and liver fatty acid-binding pro-

teins in human intestinal cell line (Caco-2): role of collagen. Exp

Cell Res 244, 441–447.

De Hoff JL, Davidson JH & Kritchevsky V (1978) An enzymatic

assay for determining free and total cholesterol in tissues. Clin

Chem 24, 433–435.

Delerive P, Furman C, Teissier E, Fruchart JC, Duriez P & Staels B

(2000) Oxidized phospholipids activate PPAR alpha in a phospho-

lipase A2-dependent manner. FEBS Lett 471, 34–38.

Deutsche Gesellschaft fur Fettwissenschaft (1994) Einheitsmethoden

zur Untersuchung von Fetten, Fettprodukten, Tensiden und ver-

wandten Stoffen. Stuttgart: Wissenschaftliche Verlagsgesellschaft.

Dietschy JM, Turley SD & Spady DK (1993) Role of liver in the

maintenance of cholesterol and low density lipoprotein homeosta-

sis in different animal species, including humans. J Lipid Res 34,

1637–1659.

Eder K (1999) The effects of a dietary oxidized oil on lipid

metabolism in rats. Lipids 34, 717–725.

S. Luci et al.880

49

Eder K, Keller U & Brandsch C (2004) Effects of a dietary oxidized

fat on cholesterol in plasma and lipoproteins and the susceptibility

of low-density lipoproteins to lipid peroxidation in guinea pigs fed

diets with different concentrations of vitamins E and C. Int J Vitam

Nutr Res 74, 11–20.

Eder K & Kirchgessner M (1998) The effect of dietary vitamin E

supply and a moderately oxidized oil on activities of hepatic

lipogenic enzymes in rats. Lipids 33, 277–283.

Eder K, Schleser S, Becker K & Korting R (2003) Conjugated lino-

leic acids lower the release of eicosanoids and nitric oxide from

human aortic endothelial cells. J Nutr 133, 4083–4089.

Endo Y, Li CM, Tagiri-Endo M & Fujimoto K (2001) A modified

method for the estimation of total carbonyl compounds in heated

and frying oils using 2-propanol as a solvent. J Am Oil Chem

Soc 78, 1021–1024.

Fan MZ, Matthews JC, Etienne NM, Stoll B, Lackeyram D & Burrin

DG (2004) Expression of apical membrane L-glutamate transpor-

ters in neonatal porcine epithelial cells along the small intestinal

crypt-villus axis. Am J Physiol Gastrointest Liver Physiol 287,

G385–G398.

Frankel EN (1998) Lipid Oxidation. Dundee: The Oily Press.

Frankel E, Smith L, Hamblin C, Creveling R & Clifford A (1984)

Occurrence of cyclic fatty acid isomers in frying fats used for

fast foods. J Am Oil Chem Soc 61, 87–90.

Guo Q, Wang PR, Milot DP, Ippolito MC, Hernandez M, Burton CA,

Wright SD & Chao Y (2001) Regulation of lipid metabolism and

gene expression by fenofibrate in hamsters. Biochim Biophys

Acta 1533, 220–232.

Habig WH, Pabst MJ & Jakoby WB (1974) Glutathione S-trans-

ferases. The first enzymatic step in mercapturic acid formation. J

Biol Chem 249, 7130–7139.

Hara A & Radin NS (1978) Lipid extraction of tissues with a low-tox-

icity solvent. Anal Biochem 90, 420–426.

Holden PR & Tugwood JD (1999) Peroxisome proliferator-activated

receptor alpha: role in rodent liver cancer and species differences. J

Mol Endocrinol 22, 1–8.

Huang C-J, Cheung N-S & Lu V-R (1988) Effects of deteriorated

frying oil and dietary protein levels on liver microsomal enzymes

in rats. J Am Oil Chem Soc 65, 1796–1803.

Keller U, Brandsch C & Eder K (2004a) The effect of dietary oxi-

dized fats on the antioxidant status of erythrocytes and their sus-

ceptibility to haemolysis in rats and guinea pigs. J Anim Physiol

Anim Nutr 88, 59–72.

Keller U, Brandsch C & Eder K (2004b) Supplementation of vitamins

C and E increases the vitamin E status but does not prevent the for-

mation of oxysterols in the liver of guinea pigs fed an oxidised fat.

Eur J Nutr 43, 353–359.

Knight BL, Hebbach A, Hauton D, Brown AM, Wiggins D & Patel

DD (2005) A role for PPARa in the control of SREBP activity

and lipid synthesis in the liver. Biochem J 389, 413–421.

Konig B & Eder K (2006) Differential action of 13-HPODE on

PPARa downstream genes in rat Fao and human HepG2 hepatoma

cell lines. J Nutr Biochem 17, 410–418.

Konig B, Koch A, Spielmann J, Hilgenfeld C, Stangl GI & Eder K

(2006) Activation of PPARa lowers synthesis and concentration

of cholesterol by reduction of nuclear SREBP-2. Biochem Pharma-

col (Epublication ahead of print version).

Kubow S (1992) Routes of formation and toxic consequences of lipid

oxidation products in foods. Free Radical Biol Med 12, 63–81.

Lemberger T, Braissant O, Juge-Aubry C, Keller H, Saladin R, Staels

B, Auwerx J, Burger AG, Meier CA & Wahli W (1996) PPAR

tissue distribution and interactions with other hormone-signaling

pathways. Ann NY Acad Sci 27, 231–251.

Lindsay CA & Wilson JD (1965) Evidence for a contribution by the

intestinal wall to the serum cholesterol in the rat. J Lipid Res 6,

173–181.

Liu JF & Huang CJ (1995) Tissue a-tocopherol retention in male rats

is compromised by feeding diets containing oxidized frying oil.

J Nutr 125, 3071–3080.

Liu JF & Huang CJ (1996) Dietary oxidized frying oil enhances tissue

a-tocopherol depletion and radioisotope tracer excretion in vitamin

E-deficient rats. J Nutr 126, 2227–2235.

Liu JF & Lee Y-W (1998) Vitamin C supplementation restores the

impaired vitamin E status of guinea pigs fed oxidised frying oil.

J Nutr 128, 116–122.

Mandard S, Muller M & Kersten S (2004) Peroxisome proliferator-

activated receptor a target genes. Cell Mol Life Sci 61, 393–416.

Marklund S & Marklund G (1974) Involvement of the superoxide

anion radical in the autoxidation of pyrogallol and a convenient

assay for superoxide dismutase. Eur J Biochem 47, 469–474.

Marrapodi M & Chiang JY (2000) Peroxisome proliferator-activated

receptor alpha (PPARalpha) and agonist inhibit cholesterol 7alpha-

hydroxylase gene (CYP7A1) transcription. J Lipid Res 41,

514–520.

Miller CW & Ntambi JM (1996) Peroxisome proliferators induce

mouse liver stearoyl-CoA desaturase 1 gene expression. Proc

Natl Acad Sci U S A 93, 9443–9448.

Mishra A, Chaudhary A & Sethi S (2004) Oxidized omega-3 fatty

acids inhibit NF-KB activation via a PPARa-dependent pathway.

Arterioscl Thromb Vasc Biol 24, 1621–1627.

Mochizuki K, Suruga K, Yagi E, Takase S & Goda T (2001) The

expression of PPAR-associated genes is modulated through post-

natal development of PPAR subtypes in the small intestine. Bio-

chim Biophys Acta 1531, 68–76.

Motojima K, Passilly P, Peters JM, Gonzalez FJ & Latruffe N (1998)

Expression of putative fatty acid transporter genes are regulated by

peroxisome proliferator-activated receptor alpha and gamma acti-

vators in a tissue- and inducer-specific manner. J Biol Chem 273,

16710–16714.

Paglia DE & Valentine WN (1967) Studies on the quantitative and

qualitative characterization of erythrocyte glutathione peroxidase.

J Lab Clin Med 70, 158–169.

Patel DD, Knight BL, Soutar AK, Gibbons GF & Wade DP (2000)

The effect of peroxisome-proliferator-activated receptor-alpha on

the activity of the cholesterol 7 alpha-hydroxylase gene. Biochem

J 351, 747–753.

Patel DD, Knight BL, Wiggins D, Humphries SM & Gibbons GF

(2001) Disturbances in the normal regulation of SREBP-sensitive

genes in PPAR alpha-deficient mice. J Lipid Res 42, 328–337.

Peffer PL, Lin X & Odle J (2005) Hepatic b-oxidation and carnitine

palmitoyltransferase I in neonatal pigs after dietary treatments of

clofibric acid, isoproterenol, and medium-chain triglycerides. Am

J Physiol Regul Integ Comp Physiol 288, R1518–R1524.

Peters JM, Cheung C & Gonzalez FJ (2005) Peroxisome proliferator-

activated receptor-alpha and liver cancer: where do we stand?

J Mol Med 83, 774–785.

Pfaffl MW (2001) A new mathematical model for relative quantifi-

cation in real-time RT-PCR. Nucleic Acids Res 29, e45.

Recknagel RO & Glende EA Jr (1984) Spectrophotometric detection

of lipid conjugated dienes. Methods Enzymol 105, 331–337.

Royall JA & Ischiropoulos H (1993) Evaluation of 20,70-dichlorofluor-

escin and dihydrorhodamine 123 as fluorescent probes for intra-

cellular H2O2 in cultured endothelial cells. Arch Biochem

Biophys 302, 348–355.

Ruiz-Gutierrez V, Perez-Espirosa A, Vazquez CM & Santa-Maria C

(1999) Effects of dietary fats (fish, olive and high-oleic-acid sun-

flower oils) on lipid composition and antioxidant enzymes in rat

liver. Br J Nutr 82, 233–241.

Schoonjans K, Staels B & Auwerx J (1996) The peroxisome prolif-

erator activated receptors (PPARs) and their effects on lipid metab-

olism and adipocyte differentiation. Biochim Biophys Acta 1302,

93–109.


50

Shakir KM, Sundaram SG & Margolis S (1978) Lipid synthesis in

isolated intestinal cells. J Lipid Res 19, 433–442.

Sidwell CG, Salwin H, Benca M & Mitchell JH Jr (1954) The use of

thiobarbituric acid as a measure of fat oxidation. J Am Oil Chem

Soc 31, 603–606.

Staprans I, Pan XM, Rapp JH & Feingold KR (2005) The role of diet-

ary oxidized cholesterol and oxidized fatty acids in the develop-

ment of atherosclerosis. Mol Nutr Food Res 49, 1075–1082.

Sulzle A, Hirche F & Eder K (2004) Thermally oxidized dietary fat

upregulates the expression of target genes of PPAR alpha in rat

liver. J Nutr 134, 1375–1383.

Walrand S, Valeix S, Rodriguez C, Ligot P, Chassagne J & Vasson

MP (2003) Flow cytometry study of polymorphonuclear neutrophil

oxidative burst: a comparison of three fluorescent probes. Clin

Chim Acta 331, 103–110.

Yoshida H & Kajimoto G (1989) Effect of dietary vitamin E on

the toxicity of autoxidized oil to rats. Ann Nutr Metab 33,

153–161.

Yu XX, Odle J & Drackley JK (2001) Differential induction of per-

oxisomal b-oxidation enzymes by clofibric acid and aspirin in

piglet tissues. Am J Physiol Regul Integr Comp Physiol 281,

R1553–R1561.

S. Luci et al.882

51

Int. J. Vitam. Nutr. Res., 76 (6), 2006, 398–406

Int. J. Vitam. Nutr. Res., 76 (6), 2006, © Hogrefe & Huber Publishers

Research PaperEffects of 13-HPODE on

Expression of Genes Involved inThyroid Hormone Synthesis,

Iodide Uptake and Formation ofHydrogen Peroxide in Porcine

ThyrocytesSebastian Luci, Anja Bettzieche, Corinna Brandsch and Klaus Eder

Institut für Ernährungswissenschaften, Martin-Luther-Universität Halle-Wittenberg, Halle/Saale, Germany

Received for publication: February 14, 2006; Accepted for publication: August 7, 2006

Abstract: It has been shown that dietary oxidized fats influence thyroid function in rats and pigs. Mechanismsunderlying this phenomenon are unknown. This study was performed to investigate whether 13-hydroperoxy-9,11-octadecadienic acid (13-HPODE), a primary oxidation product of linoleic acid, affects expression of genesinvolved in thyroid hormone synthesis and formation of hydrogen peroxide in primary porcine thyrocytes. Thy-rocytes were treated with 13-HPODE in concentrations between 20 and 100 µM. Cells treated with vehicle alone(“control cells”) or with equivalent concentrations of linoleic acid were considered as controls. Treatment ofcells with 13-HPODE did not affect cell viability but increased the activities of the antioxidant enzymes super-oxide dismutase and glutathione peroxidase (p < 0.05) compared to control cells or cells treated with linoleicacid. Relative mRNA concentrations of genes involved in thyroid hormone synthesis like sodium iodide sym-porter, thyrotropin receptor, and thyroid peroxidase, as well as iodide uptake, did not differ between cells treat-ed with 13-HPODE and control cells or cells treated with linoleic acid. Treatment of cells with 13-HPODE, how-ever, reduced the relative mRNA concentrations of dual oxidase-2 and the formation of hydrogen peroxide com-pared to control cells or cells treated with linoleic acid (p < 0.05). Because the production of hydrogen perox-ide is rate-limiting for the synthesis of thyroid hormones, it is suggested that 13-HPODE could have an impacton the formation of thyroid hormones in the thyroid gland.

Key words: 13-HPODE, thyrocytes, thyroid hormones, pig

DOI 10.1024/0300-9831.76.6.39852

S. Luci et al: Effect of 13-HPODE on Porcine Thyrocytes 399


IntroductionOxidized fats are generated during processing and storageof foods and constitute an important portion of Westerndiets. Increased intake of oxidized fats has been linked toan enhanced incidence of coronary heart disease, en-dothelial dysfunction, and cancer in humans [1, 2]. Sev-eral studies examined the physiological effects of oxidizeddietary fats in animal models [3–6]. In rats and pigs, in-creased concentrations of free and total thyroxine in plas-ma were observed after feeding a diet rich in oxidized fats[7, 8]. In rats, a dietary oxidized fat, moreover, led to al-terations in the morphology of the thyroid gland and in theexpression of genes involved in thyroid hormone synthe-sis. Rats fed oxidized fat exhibited an increased height ofepithelial cells while the diameter of follicles was reduced;gene expression of sodium iodide symporter (NIS) wasreduced and that of thyroid peroxidase (TPO) increasedin the thyroid glands of these rats [9]. These alterationssuggest that dietary oxidized fat affects the function of thethyroid gland, i.e. activities of proteins involved in the for-mation of thyroid hormones. The components responsiblefor the effects of oxidized fat and the mechanisms by whichthey influence thyroid function are unknown.

Oxidized fats include a mixture of primary and sec-ondary lipid peroxidation products depending on theirthermal treatment. Heating fats for a long period at lowtemperature without catalysts produces mainly primarylipid peroxidation products such as hydroxy or hydroper-oxy fatty acids. Since primary lipoxy radicals are unsta-ble, fats treated at high temperature or in the presence ofcatalysts contain predominantly secondary lipid peroxi-dation products such as carbonyls or dimeric, trimeric,polymeric, and cyclic fatty acids [10]. In our previous stud-ies, feeding fats that had been treated at a low temperaturecaused a stronger increase of plasma thyroxine concen-trations in rats than feeding a fat treated at a high temper-ature [7]. Therefore, we assume that primary lipid perox-idation products may be responsible for the effects of ox-idized fat on thyroid hormone metabolism. A number ofstudies have shown that lipid oxidation products are read-ily absorbed, incorporated into chylomicrons as well asvery-low (VLDL) and low-density lipoproteins (LDL),and are taken up in body cells [11–14]. 13-hydroperoxy-9,11-octadecadienic acid (13-HPODE) is the primary au-toxidation product of linoleic acid [15]. It originates notonly from the diet but is also formed in the body by radi-cal-driven non-enzymatic processes and by the action of15-lipoxygenase (15-LOX), an enzyme that occurs inmany human tissues [16]. Lipid hydroperoxides such as13-HPODE have a strong impact on the metabolism ofcells. As a component of cells, oxidized fatty acids pro-duce oxidative stress, exert cytotoxic effects [17], and, as

natural ligands of the alpha and gamma peroxisome pro-liferator activated receptors (PPARα and PPARγ) they areable to influence lipid metabolism and cell differentiationin several ways [18, 19]. Recently, it has been shown that13-HPODE activates the pro-inflammatory NF-κB path-way in vascular smooth muscle cells [20, 21]. As the quan-titatively most important lipid oxidation product in oxi-dized LDL, 13-HPODE is also involved in the process ofatherosclerosis [22]. However, potential effects of 13-HPODE in thyrocytes have not yet been studied.

The aim of this study was to investigate whether 13-HPODE affects the function of thyrocytes, i.e. metabolicsteps involved in the formation of thyroid hormones. Io-dide required for thyroid hormone synthesis enters the thy-rocyte through an active process mediated by NIS locat-ed in the basal membrane [23]. TPO synthesizes thyroidhormone residues on thyroglobulin by successively cat-alyzing the iodination of tyrosyl residues and the couplingof iodotyrosine pairs into iodothyronines. These reactionstake place on the outer side of the apical plasma mem-brane of thyrocytes in the presence of hydrogen peroxideas an electron acceptor [24]. The hydrogen peroxide-gen-erating system which constitutes the rate-limiting step ofthyroid hormone synthesis [25] is a Ca2+-dependentNADPH oxidase. This enzyme contains two integral mem-brane flavoproteins called dual oxidases (DUOX) 1 and 2[26]. Recent studies suggested that DUOX2 is the majorgenerator of hydrogen peroxide in thyrocytes, while it hasbeen questioned whether DUOX1 is involved in thyroidhormone generation [27]. The catalytic activity ofNADPH oxidase is essentially triggered by the Ca2+-phos-phatidylinositol cascade [28]. All steps of thyroid hormonesynthesis are stimulated by thyrotropin (TSH), whichbinds to the TSH-receptor and increases the expression ofNIS, TPO, and DUOX genes through the cAMP or IP3

pathway, respectively [28–30]. In this study we deter-mined the effects of 13-HPODE on gene expression ofNIS, TPO, DUOX2, and TSH-receptor, on iodide uptake,and on formation of hydrogen peroxide in primary porcinethyrocytes, used as a model of thyroid cells that express-es all important genes of thyroid hormone synthesis.

Oxidized fatty acids are able to cause oxidative stressin cells by producing superoxide anions, which may af-fect the cellular antioxidant system [31]. To study whether13-HPODE affects the antioxidant status of thyrocytes,we determined the activities of the most important cellu-lar antioxidant enzymes, namely superoxide dismutase(SOD), glutathione peroxidase (GSH-Px), and catalase.The effects of 13-HPODE were compared with those oflinoleic acid (LA) or vehicle alone.

53

400 S. Luci et al: Effect of 13-HPODE on Porcine Thyrocytes


Materials and MethodsIsolation and treatment of primary porcinethyrocytes

Thyroid glands from 10-week-old pigs used in trainingprograms of the Faculty of Medicine were transported onice and processed within one hour of death. Thyrocyteswere isolated from glands as described previously [32].Briefly, the glands were stripped of connective tissue andcut into ~2 mm pieces. These fragments were washed inHank’s buffered salt solution (HBSS) and sedimented at100 × g for 2 minutes. The pellets were resuspended andincubated in HBSS containing 2% collagenase (BiochromAG, Berlin, Germany) for 30 minutes at 37°C. The digestwas dissociated by repeated pipetting, filtered through ny-lon mesh, and pelleted at 100 × g for 1 minute. The pelletwas washed 6 times in Dulbecco’s modified Eagle medi-um (DMEM) supplemented with gentamicin (0.5%). Thefinal pellet was resuspended in DMEM supplemented withgentamicin (0.5%) and fetal calf serum (FCS) (10%), plat-ed in cell culture vessels, and incubated at 37°C in an in-cubator with a humidified atmosphere and 5% CO2 for 4days without further passage. Medium was changed after24 hours (complete medium) and 72 hours (FCS-freemedium). Thyrocytes were incubated for 2 hours or 24hours in FCS-free medium containing either 20, 40, or 100µmoles 13-HPODE or LA or ethanol as vehicle alone (con-trol cells).

Preparation of 13-HPODE

Stock solution of linoleic acid (Sigma-Aldrich GmbH,Taufkirchen, Germany) was prepared in absolute ethanol.The linoleic acid was oxidized to 13-HPODE with im-mobilized soybean lipoxygenase (100 U/mL, BiochromAG, Berlin, Germany) at 37°C for 1 hour. The formationof 13-HPODE was monitored spectrophotometrically byscanning the absorption between 200 and 300 nm usingphosphate buffered saline as a reference [33]. Under theseconditions, the conversion into 13-HPODE is observed asan increase in absorbance at an optical density of 234 nm.Usually, more than 90% conversion of linoleic acid to 13-HPODE was achieved as determined by the molar ex-tinction coefficient of conjugated dienes (E = 29,500 ×mol-1 × cm-1). 13-HPODE was extracted with n-hexane,dried under vacuum, and resuspended in ethanol. Con-centration of 13-HPODE in the alcohol stock solution waschecked by measuring the absorbance at 234 nm. Variousdilutions of the stock solutions were used for the incuba-tions.

Viability and iodide uptake

The viability of the cells was determined following treat-ment with either 13-HPODE or LA in a concentrationrange from 20 to 100 µM for 2 hours or 24 hours by MTTassay.

Iodide uptake was measured in cells seeded into 6 wellplates. After treatment of the cells with 20 µmoles of ei-ther 13-HPODE or LA or vehicle alone for 24 hours themedium was removed and 2.0 mL of HBSS containingNa125I (0.3 µCi/mL) were added to each well. For estima-tion of specific uptake by NIS, sodium perchlorate wasadded to the incubation medium at a final concentrationof 1 mM. Cells were incubated at 37°C for 5 or 30 min-utes. The incubation was terminated by aspiration of themedium. After rinsing 3 times with ice-cold Hank’sbuffered salt solution (HBSS), cells were lysed with 1 mLIgepal-lysis buffer [50 mM Tris, 140 mM NaCl, 1.5 mMMgSO4 × 7 H2O, 0.5% (v/v) Igepal, pH 8.0] and scrapedfrom each well for 125I counting.

Determination of hydrogen peroxide

The generation of hydrogen peroxide was quantified bymeasuring the oxidation of dihydrorhodamine 123 (DHR)into the fluorescent product rhodamine 123 (excitationwavelength: 485 nm, emission wavelength: 538 nm) [34].After treatment of cells seeded into 24-well plates, DHRsolution was added either to the cells with the incubationmedium for analysis of total hydrogen peroxide concen-tration or after exchange of the incubation medium byphosphate-buffered saline (PBS) for analysis of the intra-cellular hydrogen peroxide concentration. The final con-centration of DHR was 27.5 µM. The amount of hydro-gen peroxide secreted into the medium was calculated bysubtracting the intracellular from the total concentration.It has been shown that DHR is specifically oxidized byhydrogen peroxide [34]. There was no interference of thedetermination of hydrogen peroxide with 13-HPODEadded to the medium. Cell protein was assayed by theBicinchoninic Acid (BCA) protein assay method. Valuesare given as fluorescence intensity per mg cell protein.

Measurement of activities of SOD, GSH-Px,and catalase

For measurement of enzyme activities after treatment,cells were dislodged from culture vessels by scraping.Cells were pelleted, resuspended in PBS, and disintegrat-ed with ultrasound 2 times for 30 seconds each. SOD ac-tivity was determined according to the method of Mark-lund and Marklund [35] with pyrogallol as the substrate.The activity of GSH-Px was determined with t-butyl hy-

54



droperoxide at 25°C according to the method of Pagliaand Valentine [36]. Catalase activity was determined at25°C using hydrogen peroxide as substrate according tothe method of Aebi [37]. All enzyme activities were re-lated to the protein concentration of the analyzed cells.

Determination of conjugated dienes

Total lipids of cells seeded into 6-well plates were ex-tracted with 2 mL hexane. The absorbance of the extractwas measured at a wavelength of 234 nm and the con-centration of dienes was related to the protein concentra-tion of the analyzed cells. Concentration of conjugated di-enes was calculated by using the molar extinction coeffi-cient for conjugated dienes at 234 nm (E = 29,500 × mol-1

× cm-1).

Isolation of RNA and semi-quantitative RT-PCR

Total RNA from thyrocytes was isolated by using Trizolreagent (Invitrogen, Karlsruhe, Germany) according to themanufacturers protocol. The relative mRNA quantities ofNIS, TPO, TSH receptor, and DUOX2, related to the ref-erence gene glyceraldehyde-3-phosphate-dehydrogenase(GAPDH) mRNA, were determined by means of reverse-transcriptase polymerase chain reaction (RT-PCR). Firststrand cDNA synthesis was performed from 1.2 µg totalRNA by reverse transcription using the RevertAid™ M-MuLV reverse transcriptase (MBI Fermentas, St. Leon-Rot, Germany) and oligo dT primers (Operon, Köln, Ger-many). cDNA was amplified in a 20 µL reaction contain-ing 2 µL RT-mixture, 0.2 µL Biotherm™ DNA poly-merase, 2 µL 10X PCR buffer, 0.4 µL DNA polymeriza-tion mix (all from Genecraft, Lüdinghausen, Germany),and gene-specific primers obtained from Carl Roth(Karlsruhe, Germany). The primer sequences used wereas follows: 5’-AGT-CAT-CAG-CGG-CCC-CCT-CCT-C-3’ (forward) and 5’-ACC-GAT-GCC-GTC-TGC-CGT-GTG-3’ (reverse) for pig NIS; 5’-CTG-GGC-GCC-GTG-CTC-GTC-TG-3’ (forward) and 5’-ACG-CGG-GTG-GCA-TCT-GAC-TCT-GAC-3’ (reverse) for pig TPO; 5’-GCC-TGC-CCA-TGG-ACA-CTG-AGA-C-3’ (forward)and 5’-CTG-ACC-CCG-GTA-TGC-CTG-AGC-3’ (re-verse) for pig TSH receptor; 5’-GAC-CCA-GCG-GCA-GTT-TGA-ATG-G-3’(forward) and 5’-AGG-GCC-GCA-GCT-GAA-CAC-TCC-3’ (reverse) for pig DUOX2 and5’-AGG-GGC-TCT-CCA-GAA-CAT-CAT-CC-3’(forward)and 5’-TCG-CGT-GCT-CTT-GCT-GGG-GTT-GG-3’ (re-verse) for pig GAPDH.

Statistical analysis

Means of the treatments (13-HPODE, LA) and controlwere compared by Student’s t-test using the Minitab Sta-tistical Software (Minitab, State College, PA, USA). Da-ta were considered significantly different at p < 0.05.

Results

Viability and iodide uptake into thyroidcells

Viability of primary porcine thyrocytes was not influencedby treatment with LA or 13-HPODE up to concentrationsof 100 µM. When examined under the light microscope,the cells appeared normal during the whole incubation pe-riod of 24 hours. Radioiodide was taken up by the cells ina time-dependent manner and this uptake was partially in-hibited by perchlorate, indicating active and specific up-take of iodide via NIS. However, there was no differencein total iodide uptake and iodide uptake by NIS betweencontrol cells and cells treated with 20 µM of 13-HPODEor LA for 24 hours (Figure 1).

Concentration of hydrogen peroxide

Hydrogen peroxide was measurable in the cells as well asin the cell medium. Cells treated for 24 hours with 40µmoles of 13-HPODE released less (p < 0.05) hydrogen

0

2

4

6

8

10

12

14

Total, 5 min Total, 30 min NIS, 30 min

Iod

ide u

pta

ke (

10

5 c

pm

/105

cell

s)

Control LA 13-HPODE

Figure 1: Effect of 13-HPODE on uptake of 125I by primaryporcine thyrocytes. Cells were treated with 20 µmoles of 13-HPODE or linoleic acid (LA) or with vehicle alone (control) for24 hours. Thereafter, cells were incubated at 37°C for 5 or 30minutes with Na125I alone (Total) or together with sodium per-chlorate (1 mM) for determination of specific iodide uptake bysodium iodide symporter (NIS). Values are means ± SD (n = 2).

55



peroxide into the medium during the incubation than cellstreated with 40 µmoles of LA or control cells [Fluores-cence intensity (AU * 103)/mg protein: Control, 37.0 ±11.5; cells treated with 40 µmoles of LA, 28.1 ± 7.0; cellstreated with 40 µmoles of 13-HPODE, 20.0 ± 8.1, means± SD, n = 8 for each treatment]. The intracellular con-centration of hydrogen peroxide tended (p < 0.15) also tobe lower in cells treated with 13-HPODE than in cells treat-ed with LA or in control cells [Fluorescence intensity (AU* 103)/mg protein: Control, 4.6 ± 1.3; cells treated with 40µmoles of LA, 4.4 ± 0.9; cells treated with 40 µmoles of13-HPODE, 3.9 ± 1.2, means ± SD, n = 8 for each treat-ment].

Activities of antioxidant enzymes andconcentrations of conjugated dienes

Cells treated for 24 hours with 40 µmoles of 13-HPODEhad higher activities of SOD and GSH-Px than cells treat-ed with 40 µmoles of LA or control cells (p < 0.05, Fig-ure 2). The activity of catalase did not differ between cellstreated for 24 hours with 40 µmoles of LA or 13-HPODEand control cells (Figure 2). The concentration of conju-gated dienes was measured in cells treated with 40 µmolesof LA and in cells treated with 40 µmoles of 13-HPODE,for 24 hours. It was significantly higher in cells treatedwith 13-HPODE than in cells treated with LA (11.1 ± 0.8vs. 2.6 ± 0.8 µmol/mg protein, n = 3 for each treatment, p< 0.05).

Relative mRNA levels of proteins involvedin thyroid hormone synthesis

Treatments were done with 100 µmoles of LA or 13-HPODE and with vehicle alone (control cells) for 2 and24 hours. Treatment with both LA or 13-HPODE did notchange the relative mRNA level of NIS, TPO, and TSHreceptor relative to control cells treated with vehicle alone(Figure 3). Relative mRNA level of DUOX2 was notchanged by treatment with LA or 13-HPODE for 2 hours(Figure 3). However, cells treated for 24 hours with 100µmoles of 13-HPODE lowered the relative mRNA levelof DUOX2 compared with cells treated with 100 µmolesof LA or control cells (p < 0.05, Figure 3).

0

0.2

0.4

0.6

0.8

1

1.2

1.4

1.6

1.8

NIS TSH-R TPO DuOX2

rela

tive m

RN

A l

evel

Control

LA

13-HPODE

2 h

0

0.2

0.4

0.6

0.8

1

1.2

1.4

NIS TSH-R TPO DuOX2

rela

tive m

RN

A l

evel

Control

LA

13-HPODE

24 h

*

0

0.2

0.4

0.6

0.8

1

1.2

1.4

1.6

1.8

2

SOD GSH-Px Catalase

Acti

vit

y (

rela

tive)

Control

LA

13-HPODE**

Figure 2: Effect of 13-HPODE on activities of superoxide dis-mutase (SOD), glutathione peroxidase (GSH-Px), and catalasein primary porcine thyrocytes. Cells were treated with 40 µmolesof 13-HPODE or linoleic acid (LA) or with vehicle alone (con-trol) for 24 hours. Enzyme activities in the cells were measuredby spectrophotometric assays. Activities were related to the val-ues of the control group. Values are means ± SD (n = 3). *Sig-nificantly different from control cells, p < 0.05.

Figure 3: Effect of 13-HPODE on relative mRNA concentra-tions of sodium iodide symporter (NIS), thyrotropin receptor(TSH-R), thyroid peroxidase (TPO), and dual oxidase-2(DUOX2) in primary porcine thyrocytes. Cells were treated ei-ther with 100 µmoles of 13-HPODE or linoleic acid (LA) or withvehicle alone (control) for 2 hours (upper panel), or for 24 hours(lower panel). Relative mRNA concentrations were determinedby RT-PCR using GAPDH mRNA for normalization. Concen-trations were related to the values of the control group. Valuesare means ± SD (n = 4). *Significantly different from controlcells, p < 0.05.

56



DiscussionPrevious studies have shown that dietary oxidized fats in-fluence the thyroid function in rats and pigs [6–9]. In thisstudy we treated porcine thyrocytes with 13-HPODE, anoxidation product of LA. We are aware of the fact that ox-idized fats contain many products which are formed dur-ing the initial stage of lipid peroxidation such as hydrox-yl, keto, and epoxy linoleic acids as well as end-productssuch as aldehydes, which may have complex effects onthyrocyte metabolism [10]. Therefore, 13-HPODE alonesurely cannot resemble the effects of an oxidized fat. Nev-ertheless, 13-HPODE is widely used to study effects ofoxidized fatty acids on function of various cell types as itis quantitatively the most important primary oxidationproduct of LA [19–22, 38–42]. Therefore, treatment with13-HPODE should provide preliminary data on whetheroxidized fatty acids could influence the function of thy-rocytes.

In most of the studies dealing with the effects of 13-HPODE on cellular function, limited concentrations of 5to 50 µmol/L were used, as higher concentrations couldbe cytotoxic [38, 39, 42]. According to this caveat, in theinitial experiments dealing with the effects of 13-HPODEon iodide uptake, hydrogen peroxide production, and ac-tivities of antioxidant enzymes, we incubated cells with20 or 40 µmol/L. As the MTT test showed that cell func-tion is not affected even by 100 µmol/L, in the experimentsinvestigating the expression of genes involved in thyroidhormone synthesis, we used concentrations of 100 µmol/L, which can be considered as very high relative to thosecommonly used in other studies. To date there is less in-formation about the uptake and metabolism of 13-HPODEin thyrocytes. It has been shown that oxidized fatty acidsare generally poorly taken up by cells compared to unox-idized free fatty acids [38]. The mechanism of uptake ofoxidized fatty acids is different from unoxidized fatty acidsand may not involve cellular expression of CD 36. Uptakeof oxidized fatty acids depends on specific cell type, andis not known for thyrocytes [38]. In vivo, a large portionof oxidized fatty acids may enter the cell as a componentof oxidized LDL [22]. Nevertheless, in most studies cellsare treated directly with 13-HPODE rather than with ox-idized LDL because the application of complex lipid sys-tems could exert some effects on cell function that are notcaused by oxidized fatty acids but by other lipid compo-nents. Within the cell, oxidized fatty acids are poorly usedby microsomal acyltransferases and are therefore incor-porated less efficiently into cell lipids [38]. Most of thebiological effects of 13-HPODE and other oxidized fattyacids are probably caused by interaction with cell-surfacecomponents [38]. In Caco-2 cells, 13-HPODE is almostcompletely reduced by GSH-Px to 13-HODE [43]. The

fate of 13-HPODE administered to thyrocytes has not yetbeen investigated. Therefore, the possibility exists that inthyrocytes a large part of 13-HPODE was also reduced to13-HODE by the action of GSH-Px.

The current study reveals that incubation of porcine thy-rocytes with 13-HPODE increases the activities of SODand GSH-Px, and we assume that the increased activitiesof these antioxidant enzymes may be the consequence ofoxidative stress caused by 13-HPODE. It has been shownthat 13-HPODE induces oxidative stress by stimulatingthe generation of superoxide radicals or hydrogen perox-ide [18, 44], and that oxidative stress stimulates gene ex-pression and activities of SOD and GSH-Px in order toprotect the cells against reactive oxygen species [45–48].Concentrations of conjugated dienes are considered as amarker of lipid peroxidation [33]. Measurement of the ab-sorbance at 234 nm shows that cells treated with 13-HPODE had much higher concentrations of conjugateddienes than cells treated with LA, indicative of increasedabsolute concentrations of oxidized fatty acids. Increasedconcentrations of conjugated fatty acids can occur for twodifferent reasons. First, 13-HPODE and its decompositionproduct 13-HODE themselves have conjugated dienestructures, and their uptake into the cell contributes to in-creased cellular concentrations of dienes. Second, the 13-HPODE that enters the cell could further propagate oxi-dation of membrane-bound fatty acids that contribute toincreased levels of conjugated dienes. In this respect, it isalso likely that oxidized fatty acids from arachidonic acid,such as hydroxyeicosatetraenoic acids (HETEs) or hy-droperoxytetraenoic acids (HPETEs), are formed. Thesefatty acids have been reported to exert many biologic ef-fects in various cell types. For example, they act chemo-tactically and cytotoxically, induce cell proliferation, andactivate various signal transduction pathways in variouscell types [49–51]. Although no information about the ef-fects of HETEs and HPETEs on thyrocyte function isavailable, it is likely that these fatty acids could contributeto the effects observed in cells treated with 13-HPODE. Itis well known that peroxidation of lipids can be prevent-ed by antioxidants [33]. To assess whether oxidation prod-ucts of polyunsaturated fatty acids (PUFA) formed duringincubation are involved in the effects of 13-HPODE, fur-ther studies should consider the interaction between 13-HPODE and the supply of cells with antioxidants.

The present study shows that incubation of porcine thy-rocytes with 13-HPODE does not lead to alterations ingene expression of NIS and TPO or iodide uptake, evenin nonphysiologic high concentrations of 100 µM over arelatively long period of 24 hours. Gene expression of NISand TPO is primarily regulated by TSH. Binding of TSHto the TSH receptor leads to the release of cAMP, whichin turn enhances gene expression of NIS and TPO [29, 30].

57



The observation that gene expression of the TSH receptorwas also not influenced by 13-HPODE suggests that it al-so did not influence the effect of TSH on the function ofporcine thyrocytes. Gene expression of NIS, however, isnot only regulated by TSH through the cAMP pathway butalso by tumor necrosis factor-alpha (TNFα), a cytokinewhose release is stimulated by activation of NF-κB [52,53]. Recently, it has been shown that 13-HPODE leads toan activation of NF-κB in vascular smooth muscle cells[20, 21]. It has been observed that NF-κB activity is alsopresent in thyrocytes [54, 55]. The results of the presentstudy indirectly suggest that 13-HPODE did not activateNF-κB in porcine thyrocytes, because an activation of thisnuclear factor would have led to a down-regulation of NISexpression.

The present study shows that high concentrations of 13-HPODE lead to a down-regulation of the DUOX2 gene andto a reduced release of hydrogen peroxide. We did not mea-sure the activity of NADPH oxidase but we assume that areduced gene expression of DUOX2 might have been as-sociated with a reduced activity of this enzyme. The mech-anism by which high concentrations of 13-HPODE reducegene expression of DUOX2 remains to be elucidated.

The reduced concentrations of hydrogen peroxide inthe cells and in the cell medium could be due to the in-creased activity of GSH-Px observed in cells treated with13-HPODE. GSH-Px protects thyrocytes against a highintracellular concentration of hydrogen peroxide, whichfor instance can lead to apoptosis [56]. It has been postu-lated that GSH-Px in thyrocytes acts as a regulator of thy-roid hormone biosynthesis by controlling the concentra-tion of hydrogen peroxide available for thyroid hormonesynthesis [57].

As the concentration of hydrogen peroxide in thyro-cytes is the rate-limiting factor of thyroid hormone syn-thesis [25], it can be suggested that high concentrations of13-HPODE could lead to a reduced formation of thyroidhormones. Recently, it has been shown that generation ofreactive oxygen species inhibit the formation of thyroidhormones in cultured thyroid cells [58]. It cannot be ex-cluded that the effects observed on DUOX2 expressionand release of hydrogen peroxide by 13-HPODE are due,at least in part, to reactive oxygen species produced or tolipid oxidation products formed during incubation in thecell. It is possible that a reduction of the release of hy-drogen peroxide could lead to a reduced formation of thy-roid hormones in thyrocytes. Whether 13-HPODE has aneffect on the release of hydrogen peroxide in the thyroidand the formation of thyroid hormones in vivo remains tobe elucidated.

Previously, we have observed that feeding a dietary ox-idized fat leads to a reduced gene expression of NIS andan increased gene expression of TPO in the thyroid gland

and an increased concentration of thyroxine in the bloodof rats [7, 9]. The present study reveals that these effectsprobably are not caused by 13-HPODE, the quantitative-ly most important oxidation product of LA, or by othersecondary lipid oxidation products that are formed in thecell during incubation with 13-HPODE.

In conclusion, this study shows that incubation ofporcine thyrocytes with 13-HPODE does not change geneexpression of NIS, TPO, and TSH receptor, or iodide up-take into the cell. High concentrations of 13-HPODE,however, reduced gene expression of DUOX2 and pro-duction of hydrogen peroxide. Because the production ofhydrogen peroxide is rate-limiting for the synthesis of thy-roid hormones, it cannot be excluded that oxidized fattyacids could have an impact on the formation of thyroidhormones in the thyroid gland.

Acknowledgment

The authors would like to thank J. Thielebein for prepa-ration of thyroid glands from the pigs.

References1. Cohn, J.S. (2002) Oxidized fat in the diet, postprandial li-

paemia and cardiovascular disease. Curr. Opin. Lipidol. 13,19–24.

2. Yang, C.M., Kendall, C.W., Stamp, D., Medline, A., Archer,M.C. and Bruce, W.R. (1998) Thermally oxidized dietaryfat and colon carcinogenesis in rodents. Nutr. Cancer 30,69–73.

3. Corcos Benedetti, P., D’Aquino, M., Di Felice, M., Gentili,V., Tagliamonte, B. and Tomassi, G. (1987) Effects of a frac-tion of thermally oxidized soy bean oil on growing rats. Nu-tr. Rep. Int. 36, 387–401.

4. Blanc, P., Revol, A. and Pacheco, H. (1992) Chronic inges-tion of oxidized oil in the rat: effect on lipid composition andon cytidylyl transferase activity in various tissues. Nutr. Res.12, 833–844.

5. Hochgraf, E., Mokady, S. and Cogan, U. (1997) Dietary ox-idized linoleic acid modifies lipid composition of rat livermicrosomes and increases their fluidity. J. Nutr. 127, 681–686.

6. Eder, K., Suelzle, A., Skufca, P., Brandsch, C. and Hirche,F. (2003) Effects of dietary thermoxidized fats on expressionand activities of hepatic lipogenic enzymes in rats. Lipids38, 31–38.

7. Eder, K., Skufca, P. and Brandsch, C. (2002) Thermallyoxidized dietary fats increase plasma thyroxine concentra-tions in rats irrespective of the vitamin E and selenium sup-ply. J. Nutr. 132, 1275–1281.

8. Eder, K. and Stangl, G. I. (2000) Plasma thyroxine andcholesterol concentrations of miniature pigs are influencedby thermally oxidized dietary lipids. J. Nutr. 130, 116–121.

58



9. Skufca, P., Brandsch, C., Hirche, F. and Eder, K. (2003) Theeffects of a dietary thermally oxidized fat on thyroid mor-phology and mRNA concentrations of thyroidal iodide trans-porter and thyroid peroxidase in rats. Ann. Nutr. Metab. 47,207–213.

10. Chang, S.S., Peterson, R. J. and Ho, C.T. (1978) Chemicalreactions involved in the deep-fat frying of foods. J. Am. OilChem. Soc. 55, 718–727.

11. Staprans, I., Pan, X.M., Miller, M. and Rapp, J.H. (1993)Effect of dietary lipid peroxides on metabolism of serumchylomicrons in rats. Am. J. Physiol. 264, G561–G568.

12. Staprans, I., Rapp, J.H., Pan, X.M. and Feingold, K.R.(1993) The effects of oxidized lipids in the diet on serumlipoprotein peroxides in control and diabetic rats. J. Clin. In-vest. 92, 638–643.

13. Staprans, I., Rapp, J.H., Pan, X.M., Hardman, D.A. andFeingold, K.R. (1996) Oxidized lipids in the diet acceleratethe development of fatty streaks in cholesterol-fed rabbits.Arterioscler. Thromb. Vasc. Biol. 16, 533–538.

14. Penumetcha, M., Khan, N. and Parthasarathy, S. (2000) Di-etary oxidized fatty acids: an atherogenic risk? J. Lipid Res.41, 1473–1480.

15. Niki, E., Yoshida, Y., Saito, Y. and Noguchi, N. (2005) Lipidperoxidation: Mechanism, inhibition, and biologic effects.Biochem. Biophys. Res. Commun. 336, 1–9.

16. Kuhn, H. and Borngraber, S. (1999) Mammalian 15-lipoxy-genases. Enzymatic properties and biological implications.Adv. Exp. Med. Biol. 447, 5–28.

17. Li, W.G., Stoll, L.L., Rice, J.B., Xu, S.P., Miller, F. J. Jr,Chatterjee, P., Hu, L., Oberley, L.W., Spector, A.A. and Wein-traub, N.L. (2003) Activation of NAD(P)H oxidase by lipidhydroperoxides: mechanism of oxidant-mediated smoothmuscle cytotoxicity. Free Rad. Biol. Med. 34, 937–946.

18. Mishra, A., Chaudhary, A. and Sethi, S. (2004) Oxidizedomega-3 fatty acids inhibit NF-KB activation via a PPARα-dependent pathway. Arterioscl. Thromb. Vasc. Biol. 24,1621–1627.

19. König, B. and Eder, K. (2006) Differential action of 13-HPODE on PPARα downstream genes in rat Fao and humanHepG2 hepatoma cell lines. J. Nutr. Biochem. 17. 410–418.

20. Natarajan, R., Reddy, M.A., Malik, K.U., Fatima, S. andKhan, B.V. (2001) Signalling mechanism of Nuclear Fac-tor-kB-mediated activation of inflammatory genes by 13-hy-droperoxyoctadecadienoic acid in cultured vascular smoothmuscle cells. Arterioscler. Thromb. Vasc. Biol. 21, 1408–1413.

21. Dwarakanath, R.S., Sahar, S., Reddy, M.A., Castanotto, D.,Rossi, J. J. and Natarajan, R. (2004) Regulation of monocytechemoattractant protein-1 by the oxidized lipid, 13-hy-droperoxyoctadecadienoic acid, in vascular smooth musclecells via nuclear factor-kappa B (NF-κB). J. Molecul. Cell.Cardiol. 36, 585–595.

22. Folcik, V.A. and Cathcart, M.K. (1994) Predominance ofesterified hydroperoxy-linoleic acid in human monocyte-ox-idized LDL. J. Lipid Res. 35, 1570–1582.

23. Dai, G., Levy, O. and Carrasco, N. (1996) Cloning and char-acterization of the thyroid iodide transporter. Nature 379,458–460.

24. Ekholm, R. (1981) Iodination of thyroglobulin. An intracel-lular or an extracellular process? Mol. Cell Endocrinol. 24,141–163.

25. Corvilain, B., Van Sande, J., Laurent, E. and Dumont, J.E.(1991) The H2O2-generating system modulates protein iod-ination and the activity of pentose phosphate pathway in dogthyroid. Endocrinology 128, 779–785.

26. De Deken, X., Wang, D., Many, M.C., Costagliola, S., Lib-ert, F., Vassart, G., Dumont, J.E. and Miot, F. (2000) Cloningof two human thyroid cDNAs encoding new members of theNADPH oxidase family. J. Biol. Chem. 275, 23227–23233.

27. Moreno, J.C., Bikker, H., Kempers, M. J., van Trotsenburg,A.S., Baas, F., de Vijlder, J. J., Vulsma, T. and Ris-Stalpers,C. (2002) Inactivating mutations in the gene for thyroid ox-idase 2 (THOX2) and congenital hypothyroidism. N. Engl.J. Med. 347, 95–102.

28. Raspe, E. and Dumont, J.E. (1995) Tonic modulation of dogthyrocyte H2O2 generation and I- uptake by thyrotropinthrough the cyclic adenosine 3’, 5’-monophosphate cascade.Endocrinol. 136, 965–973.

29. Dohan, O., De la Vieja, A., Paroder, V., Riedel, C., Artani,M., Reed, M., Ginter, C.S. and Carrasco, N. (2003) The sodi-um/iodide symporter (NIS): characterization, regulation,and medical significance. Endocrine Rev. 24, 48–77.

30. McLachlan, S.M and Rapoport, B. (1992) The molecularbiology of thyroid peroxidase: cloning, expression and roleas autoantigen in autoimmune thyroid disease. EndocrineRev. 13, 192–206.

31. Wang, T.-G., Gotoh, Y., Jennings, M.H., Rhoads, C.A. andAw, T.Y. (2000) Lipid-hydroperoxide-induced apoptosis inhuman colonic CaCo-2 cells is associated with early loss ofcellular redox balance. FASEB J. 14, 1567–1576.

32. Pocar, P., Klonisch, T., Brandsch, C., Eder, K., Fröhlich, C.,Hoang-Vu, C. and Hombach-Klonisch, S. (2006) AhR-Ag-onist-Induced Transcriptional Changes of Genes Involved inThyroid Function in Primary Porcine Thyrocytes. Toxicol.Sci. 89, 408–414.

33. Esterbauer, H., Gebicki, J., Puhl, H. and Jürgens, G. (1992)The role of lipid peroxidation and antioxidants in oxidativemodification of LDL. Free Rad. Biol. Med. 13, 341–390.

34. Walrand, S., Valeix, S., Rodriguez, C., Ligot, P., Chassagne,J. and Vasson, M.P. (2003) Flow cytometry study of poly-morphonuclear neutrophil oxidative burst: a comparison ofthree fluorescent probes. Clin. Chim. Acta, 331, 103–110.

35. Marklund, S. and Marklund, G. (1974) Involvement of thesuperoxide anion radical in the autoxidation of pyrogalloland a convenient assay for superoxide dismutase. Eur. J.Biochem. 47, 469–474.

36. Paglia, D.E. and Valentine, W.N. (1967) Studies on the quan-titative and qualitative characterization of erythrocyte glu-tathione peroxidase. J. Lab. Clin. Med. 70, 158–160.

37. Aebi, H.E. (1986) Catalase. In: Methods of enzymatic analy-sis (Bergmeyer, H.U., ed.), pp. 273–286, Third Edition, Band3, VCH Verlagsgesellschaft mbH, Weinheim.

38. Auge, N., Santanam, N., Mori, N.M., Keshava, C. andParthasarathy, S. (1999) Uptake of 13-hydroperoxylinoleicacid by cultured cells. Arterioscler. Thromb. Vasc. Biol. 19,925–931.

59



39. Glasgow, W.C., Hui, R., Everhart, A.L., Jayawickreme, S.P.,Angerman-Stewart, J., Han, B.B. and Eling, T.E. (1997) Thelinoleic acid metabolite, (13S)-hydroperoxyoctadecadienoicacid, augments the epidermal growth factor receptor signal-ing pathway by attenuation of receptor dephosphorylation.Differential response in Syrian hamster embryo tumor sup-pressor phenotypes. J. Biol. Chem. 31, 19269–19276.

40. Friedrichs, B., Toborek, M., Hennig, B., Heinevetter, L.,Müller, C. and Brigelius-Flohe, R. (1999) 13-HPODE and13-HODE modulate cytokine-induced expression of en-dothelial cell adhesion molecules differently. Biofactors 9,61–72.

41. Hui, R., Kameda, H., Risinger, J. I., Angerman-Stewart, J.,Han, B., Barrett, J.C., Eling, T.E. and Glasgow, W.C. (1999)The linoleic acid metabolite, 13-HPODE augments the phos-phorylation of EGF receptor and SHP-2 leading to their in-creased association. Prostaglandins Leukot. Essent. FattyAcids 61, 137–143.

42. Meilhac, O., Zhou, M., Santanam, N. and Parthasarathy, S.(2000) Lipid peroxides induce expression of catalase in cul-tured vascular cells. J. Lipid Res. 41, 1205–1213.

43. Müller, C., Friedrichs, B., Wingler, K. and Brigelius-Flohe,R. (2002) Perturbation of lipid metabolism by linoleic acidhydroperoxide in CaCo-2 cells. Biol. Chem. 383, 637–648.

44. Santanam, N., Auge, N., Zhou, M., Keshava, C. andParthasarathy, S. (1999) Overexpression of human catalasegene decreases oxidized lipid-induced cytotoxicity in vas-cular smooth muscle cells. Arterioscler. Thromb. Vasc. Bi-ol. 19, 1912–17.

45. Atalay, M., Laaksonen, D.E., Khanna, S., Kaliste, K.E.,Hanninen, O. and Sen, C.K. (2000) Vitamin E regulateschanges in tissue antioxidants induced by fish oil and exer-cise. Med. Sci. Sports Exerc. 32, 601–607.

46. Reddy, A. and Fernandes, G. (1999) Modulation of antiox-idant enzymes and apoptosis in mice by dietary lipids andtreadmill exercise. J. Clin. Immunol. 19, 35–44.

47. Ruiz-Gutierrez, V., Perez-Espirosa, A., Vazquez, C.M. andSanta-Maria, C. (1999) Effects of dietary fats (fish, olive andhigh-oleic-acid sunflower oils) on lipid composition and an-tioxidant enzymes in rat liver. Br. J. Nutr. 82, 233–241.

48. Hsu, H.C., Lee, Y.T. and Chen, M.F. (2001) Effects of fishoil and vitamin E on the antioxidant defence system in diet-induced hypercholesterolemic rabbits. Prostaglandins andOther Lipid Mediators 66, 99–108.

49. Nakao, J., Ooyama, T., Ito, H., Chang, W.C. and Murota, S.(1982) Comparative effect of lipoxygenase products ofarachidonic acid on rat aortic smooth muscle cell migration.Atherosclerosis 44, 339–343.

50. Nishio, E. and Watanabe, Y. (1997) Role of the lipoxygenasepathway in phenylephrine-induced vascular smooth musclecell proliferation and migration. Eur. J. Pharmacol. 336,267–273.

51. Khan, B.V., Parthasarathy, S.S., Alexander, R.W. and Med-ford R.M. (1995) Modified low density lipoprotein and itsconstituents augment cytokine-activated vascular cell adhe-sion molecule-1 gene expression in human vascular en-dothelial cells. J. Clin. Invest. 95, 1262–1270.

52. Ajjan, R.A., Watson, P.F., Findlay, C., Metcalfe, R.A., Crisp,M., Ludgate, M. and Weetman, A.P. (1998) The sodium io-dide symporter gene and its regulation by cytokines foundin autoimmunity. J. Endocrinol. 158, 351–358.

53. Pekary, A.E. and Hershman, J.M. (1998) Tumor necrosisfactor, ceramide, transforming growth factor-beta1, and ag-ing reduce Na+/I- symporter messenger ribonucleic acid lev-els in FRTL-5 cells. Endocrinol. 139, 703–712.

54. Kikumori, T., Kambe, F., Nagaya, T., Funahashi, H. and Seo,H. (2001) Thyrotropin modifies activation of nuclear factorκB by tumour necrosis factor alpha in rat thyroid cell line.Biochem. J. 354, 573–579.

55. Kikumori, T., Kambe, F., Nagaya, T., Imai, T., Funahashi,H. and Seo, H. (2001) Activation of transcriptionally activenuclear factor-κB by tumor necrosis factor-α and its inhibi-tion by antioxidants in rat thyroid FRTL-5 cells. Endocrinol.139, 1715–1722.

56. Demelash, A., Karlsson, J.-O., Nilsson, M. and Björkmann,U. (2004) Selenium has a protective role in caspase-3-de-pendent apoptosis induced by H2O2 in primary cultured pigthyrocytes. Eur. J. Endocrinol. 150, 841–849.

57. Howie, A.F., Walker, S.W., Akesson, B., Arthur, J.R. andBeckett, G.J. (1995) Thyroidal extracellular glutathione per-oxidase: a potential regulator of thyroid-hormone synthesis.Biochem. J. 308, 713–717.

58. Sugawara, M., Sugawara, Y., Wen, K. and Giulivi, C. (2002)Generation of oxygen free radicals in thyroid cells and in-hibition of thyroid peroxidase. Exp. Biol. Med. 227,141–146.

Prof. Dr. Klaus Eder

Institut für Agrar und ErnährungswissenschaftenMartin-Luther-Universität Halle-WittenbergEmil-Abderhalden-Straße 26D-06108 Halle/Saale, GermanyFax (49) 345/5527124E-mail: [email protected]

60

PPARa agonists up-regulate organic cation transporters in rat liver cells

Sebastian Luci, Stefanie Geissler, Bettina Konig, Alexander Koch, Gabriele I. Stangl,Frank Hirche, Klaus Eder *

Institute of Agricultural and Nutritional Sciences, Martin-Luther-University Halle-Wittenberg, Emil-Abderhalden-Strasse 26,

D-06108 Halle (Saale), Germany

Received 15 September 2006Available online 27 September 2006

Abstract

It has been shown that clofibrate treatment increases the carnitine concentration in the liver of rats. However, the molecular mech-anism is still unknown. In this study, we observed for the first time that treatment of rats with the peroxisome proliferator activatedreceptor (PPAR)-a agonist clofibrate increases hepatic mRNA concentrations of organic cation transporters (OCTNs)-1 and -2 whichact as transporters of carnitine into the cell. In rat hepatoma (Fao) cells, treatment with WY-14,643 also increased the mRNA concen-tration of OCTN-2. mRNA concentrations of enzymes involved in carnitine biosynthesis were not altered by treatment with the PPARaagonists in livers of rats and in Fao cells. We conclude that PPARa agonists increase carnitine concentrations in livers of rats and cells byan increased uptake of carnitine into the cell but not by an increased carnitine biosynthesis.� 2006 Elsevier Inc. All rights reserved.

Keywords: Carnitine; Peroxisome proliferator activated receptor a; Rat; Organic cation transporter

Carnitine (L-3-hydroxy-4-N-N-N-trimethylaminobuty-rate) is an essential metabolite, which has a number ofindispensable functions in intermediary metabolism. Themost prominent function lies in its role in the transportof activated long-chain fatty acids from the cytosol to themitochondrial matrix where b-oxidation takes place. Otherfunctions of carnitine include the transfer of products ofperoxisomal b-oxidation to the mitochondria for oxidationin the citrate cycle, the modulation of the acyl-CoA/CoA-ratio, and the storage of energy as acetylcarnitine [1–3].

All tissues that use fatty acids as a fuel source requirecarnitine for normal function. Carnitine is derived fromdietary sources and endogenous biosynthesis [4]. Carnitinebiosynthesis involves a complex series of reactions involv-ing several tissues [5]. Lysine provides the carbon backboneof carnitine. Lysine in protein peptide linkages undergoesmethylation of the e-amino group to yield trimethyllysine(TML), which is released upon protein degradation.

Muscle is the major source of TML. The released TMLis further oxidized to butyrobetaine by the action of tri-methyllysine dioxygenase (TMLD), 3-hydroxy-N-TMLaldolase, and 4-N-trimethylaminobutyraldehyde dehydro-genase (TMABA-DH). Butyrobetaine is hydroxylated byc-butyrobetaine dioxygenase (BBD) to form carnitine.The last reaction which is rate-limiting for carnitine synthe-sis occurs primarily in liver and kidney [6].

Distribution within the body and intracellular homeosta-sis of carnitine are controlled by membrane transporters.The organic cation transporters (OCTNs), in particularOCTN-2, physiologically the most important one, operateon the intestinal absorption and renal reabsorption of car-nitine and play a major role in tissue distribution by catalyz-ing the uptake of carnitine into body cells. In most tissues,carnitine concentrations are much higher than in plasma,and the high tissue-to-plasma concentrations (up to about100:1 in muscle) are maintained by carnitine transporters[7]. The fact that inborn or acquired defects of OCTNs leadto primary or secondary systemic carnitine deficiency dem-onstrates their essential role in carnitine homeostasis [8].

0006-291X/$ - see front matter � 2006 Elsevier Inc. All rights reserved.

doi:10.1016/j.bbrc.2006.09.099

* Corresponding author. Fax: +345 55 27124.E-mail address: [email protected] (K. Eder).

www.elsevier.com/locate/ybbrc

Biochemical and Biophysical Research Communications 350 (2006) 704–708

BBRC

61

mailto:[email protected]

It has been shown that treatment of rats with clofibrateincreases the concentration of carnitine in the liver [9],and it was suggested that this effect is caused by increasedhepatic carnitine synthesis due to an increased availabilityof TML from muscle [10]. Clofibrate belongs to a groupof hypolipidemic compounds which exert their effects byactivation of peroxisome proliferator activated receptor(PPAR)-a, a transcription factor belonging to the nuclearhormone receptor superfamily [11]. Activation of PPARacauses an up-regulation of carnitine palmitoyltransferase(CPT)-1 and CPT-2 in the liver to enhance b-oxidation offatty acids [12,13]. Since carnitine is a cofactor of theseenzymes, activation of PPARa should increase the needfor carnitine which could be met either by an increased denovo carnitine synthesis in the liver or by an increaseduptake of carnitine from blood into the liver by membranecarnitine transporters, OCTN-1 and OCTN-2 [14]. The pos-sibility that the clofibrate induced increase in hepatic carni-tine concentration could have been mediated by anactivation of PPARa has not yet been investigated. Wehypothesized that activation of PPARa causes eitherup-regulation of enzymes involved in hepatic carnitine bio-synthesis or increases carnitine uptake into liver cells by anup-regulation of OCTNs. The hypothesis that activation ofPPARa could be involved in clofibrate induced increase ofhepatic carnitine concentration is supported by the findingthat hepatic carnitine concentration is also increased duringstarvation [15,16], a state in which PPARa is activated byincreased hepatic concentrations of unesterified fatty acids[17]. In order to investigate this hypothesis, we performedexperiments with rats and Fao rat hepatoma cells. In thefirst experiment, we treated rats with the synthetic PPARaagonist clofibrate and determined hepatic mRNA concen-trations of the carnitine transporters OCTN-1 and -2, andenzymes involved in carnitine biosynthesis (TMLD,TMABA-DH, BBD). In order to explore whether effectsof PPARa agonists on carnitine concentration are depen-dent on other tissues (e.g., muscle which provides TMLfor hepatic carnitine biosynthesis) or not, the second exper-iment was performed with rat hepatoma Fao cells whichwere treated with WY-14,643, another synthetic PPARaagonist.


Animal experiment. Male Sprague–Dawley rats, with an average initialbody weight of 366 g (±28; SD), were randomly assigned to two groups(n = 8) and kept individually in Macrolon cages in a room controlled fortemperature (22 ± 2 �C), relative humidity (50–60%), and light (12 h light/dark cycle). All experimental procedures described followed establishedguidelines for the care and handling of laboratory animals and wereapproved by the council of Saxony-Anhalt. The animals were treated with250 mg/kg of clofibrate (Fluka Chemie GmbH, Buchs, Switzerland) in1 mL sunflower oil or with an equal volume of the vehicle sunflower oil bygavage once a day 2 h after beginning of the light cycle. All rats were fed acommercial standard basal diet (‘‘altromin 1324’’, Altromin GmbH, Lage,Germany). To standardize food intake, the diets were fed daily inrestricted amounts of 18 g per day. Water was available ad libitum fromnipple drinkers during the whole experiment. At day 4 of treatment,

animals received the last dose of clofibrate or vehicle alone and 9 g of thediet and were killed 4 h later by decapitation under light anesthesia withdiethyl ether. Blood was collected into heparinized polyethylene tubes.Liver and gastrocnemius muscles were quickly removed, frozen with liquidnitrogen, and stored at �80 �C pending further analysis. Plasma wasobtained by centrifugation of the blood (1100g, 10 min, 4 �C) and storedat �20 �C.

Cell culture experiment. Fao rat hepatoma cells (ECACC, Salisbury,UK) were cultured in Ham-F12 medium supplemented with 10% FCSand 0.05 mg/mL gentamycin (Invitrogen, Karlsruhe, Germany). Cellswere maintained at 37 �C in a humidified atmosphere of 95% air and 5%CO2. For experiments, Fao cells were seeded in 6-well culture plates at adensity of 1.06 · 106 cells per well and used prior reaching confluence(usually 3 days after seeding). The cells were then stimulated for 6 and 20 hwith 50 lM WY-14,643 (Sigma–Aldrich, Steinheim, Germany). WY-14,643 was added to the medium from a stock solution in DMSO. FinalDMSO concentration did not exceed 0.1% (v/v). Cells treated with theappropriate vehicle concentration were used as a control. Cell viabilityafter treatment with WY-14,643 was assessed by (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) assay (MTT) [18]. Cell viability ofFao cells was not reduced by treatment with WY-14,643 (data not shown).

Carnitine analysis. Concentrations of free carnitine and acetyl carnitinein rat liver, muscle, plasma, and Fao cells were analyzed using tandemmass spectrometry according to Vernez et al. [19]. Quantitative analysiswas achieved by use of stable isotope-labeled internal standard carnitine-d3 (Larodan Fine Chemicals, Malmo, Sweden). Plasma or cells (50 lL)were added with methanol containing the internal standard; freeze-driedliver and muscle tissues (50 mg) were added with a water:methanol mix-ture (2:1, v/v) containing the internal standard. For extraction, the spec-imen was first sonified in an ultrasound bath for 20 min, then shaken in awater bath for 30 min at a temperature of 50 �C and finally centrifuged at13,000g for 10 min at 4 �C. The supernatant was used for further analysis.A 1100-er series HPLC (Agilent Technologies, Waldbronn, Germany)equipped with a Kromasil 100 column (5 lm particle size, 125 mm length,2 mm internal diameter, CS-Chromatographie Service, Langerwehe,Germany) and an API 2000 LC-MS/MS-System (Applied Biosystems,Darmstadt, Germany) were used for quantification of free carnitine andacetyl carnitine. For detection, the analytes were ionized by positive ion(5500 V) electrospray. As eluents, methanol and a methanol:water:aceto-nitrile mixture (50:45:5) were used.

RT-PCR analysis. Total RNA was isolated from Fao cells and ratlivers, respectively, by TRIZOL� reagent (Sigma–Aldrich, Steinheim,Germany) according to the manufacturer’s protocol. cDNA synthesis wascarried out as described [20]. The mRNA concentration of genes wasmeasured by realtime detection PCR using SYBR� Green I and a MJResearch Opticon system (Biozym Diagnostik GmbH, Oldendorf, Ger-many). Realtime detection PCR was performed with 1.25 U Taq DNApolymerase (Promega, Mannheim, Germany), 500 lM dNTPs and26.7 pmol of the specific primers (Operon Biotechnologies, Cologne,Germany; Table 1). Annealing temperature for all primers was 60 �C.Amplification efficiencies for all primer pairs were determined by templatedilution series. Calculation of the relative mRNA concentration was madeusing the amplification efficiencies and the Ct values [21]. The house-keeping gene b-actin was used for normalization.

Statistical analysis. Means of treatments and control were comparedby Student’s t test using the Minitab Statistical Software (Minitab, StateCollege, PA, USA). Differences with P < 0.05 were considered to besignificant.

Results

Carnitine concentrations in rat liver, plasma, and muscle and

in Fao cells

Rats treated with clofibrate had a higher concentrationof free carnitine in the liver than control rats (P < 0.05,

S. Luci et al. / Biochemical and Biophysical Research Communications 350 (2006) 704–708 705

62

Table 2). The concentration of acetyl carnitine which madeup however only a very small percentage of total carnitinewas lower in rats treated with clofibrate than in control rats(P < 0.05, Table 2). Concentrations of free and acetyl car-nitine in plasma were both lower in rats treated with clofi-brate than in control rats (P < 0.05, Table 2). Ingastrocnemius muscle, concentrations of free carnitinedid not differ between both groups of rats whereas the con-centration of acetyl carnitine was lower in rats treated withclofibrate than in control rats (P < 0.05, Table 2).

Fao cells treated with 50 lM WY-14,643 for either 6 or20 h had higher concentrations of free carnitine than con-trol cells treated with medium alone (P < 0.05, Table 3).Moreover, cells treated with WY-14,643 for 20 h had ahigher concentration of acetyl carnitine than control cells(P < 0.05, Table 3). After 6 h incubation, the concentration

of acetyl carnitine did not differ between cells treated withWY-14,643 and control cells (Table 3).

Relative mRNA concentrations of CPTs, OCTNs, and

enzymes involved in carnitine biosynthesis in rat liver and

Fao cells

Rats treated with clofibrate had higher relative mRNAconcentrations of CPT-1, CPT-2, OCTN-1, and OCTN-2in the liver than control rats (P < 0.05) whereas relativemRNA concentrations of genes encoding enzymes ofhepatic carnitine synthesis (TMLD, TMABA-DH, BBD)did not differ between both groups of rats (Fig. 1).

Fao cells treated with 50 lM WY-14,643 for either 6 or20 h had also higher relative mRNA concentrations ofCPT-1, CPT-2, and OCTN-2 than control cells(P < 0.05, Fig. 2). The extent of up-regulation of expres-sion of these genes by WY-14,643 was similar in cellstreated for 6 h and those treated for 20 h, indicating thata period of 6 h was already sufficient for maximum up-regulation. Relative mRNA concentrations of OCTN-1and of genes encoding enzymes of hepatic carnitinesynthesis (TMLD, TMABA-DH, BBD) did not differ

Table 1Characteristics of the specific primers used for RT-PCR analysis

Gene Forward primer (from 50 to 30) Reverse primer (from 5 0 to 30) bp NCBI GenBank

b-Actin ATCGTGCGTGACATTAAAGAGAAG GGACAGTGAGGCCAGGATACAG 429 BC063166CPT-1 GGAGACAGACACCATCCAACATA AGGTGATGGACTTGTCAAACC 416 NM_031559CPT-2 TCCTCGATCAAGATGGGAAC GATCCTTCATCGGGAAGTCA 237 NM_012930OCTN-1 AGCATTTGTCCTGGGAACAG ACTCAGGGATGAACCACCAG 200 NM_022270OCTN-2 CCTCTCTGGCCTGATTGAAG CTCCGCTGTGAAGACGTACA 226 NM_012930TMLD GCCCTGTGGCATTCAAGTAT GGTCCAACCCCTATCATGTG 201 AF374406TMABA-DH TTTGAGACTGAAGCCGAGGT CACCGGGCTGACGTTATAGT 156 NM_022273BBD ATTCTGCAAAAGCTCGGAAA CTCCTTGGAGTCCTGCTCTG 183 NM_022629

Table 2Concentrations of total carnitine in liver, plasma, and gastrocnemiusmuscle of control rats and rats treated with clofibrate

Control Clofibrate

Liver

Free carnitine (nmol/g) 282 ± 39 900 ± 145*

Acetyl carnitine (nmol/g) 12 ± 6 3 ± 1*

Plasma

Free carnitine (lmol/L) 55 ± 8 28 ± 4*

Acetyl carnitine (lmol/L) 20 ± 4 8 ± 2*

Gastrocnemius

Free carnitine (nmol/g) 631 ± 71 637 ± 63Acetyl carnitine (nmol/g) 208 ± 52 144 ± 36*

Values are means ± SD (n = 8). An asterisk (*) indicates a significantdifference from control rats (P < 0.05).

Table 3Concentrations of free carnitine and acetyl carnitine in Fao cells treatedeither with vehicle alone (control) or with WY-14,643 for 6 or 20 h

Control WY-14,643

Six hours incubation

Free carnitine (pmol/mg protein) 23 ± 8 33 ± 7*

Acetyl carnitine (pmol/mg protein) 87 ± 20 81 ± 17

Twenty hours incubation

Free carnitine (pmol/mg protein) 40 ± 7 63 ± 25*

Acetyl carnitine (pmol/mg protein) 78 ± 20 97 ± 30*

Values are means ± SD (n = 3). An asterisk (*) indicates a significantdifference from control cells (P < 0.05).

Fig. 1. Effect of clofibrate on the relative mRNA concentrations ofcarnitine palmitoyltransferases (CPT-1, CPT-2), organic cation trans-porters (OCTN-1, OCTN-2), and enzymes of carnitine biosynthesis(trimethyllysine dioxygenase, TMLD; 4-N-trimethylaminobutyraldehydedehydrogenase, TMABA-DH; c-butyrobetaine dioxygenase, BBD) in theliver of rats. Rats were treated orally with 250 mg/kg of clofibrate for 4days. Control animals obtained the appropriate volume of the vehiclesunflower oil. Total RNA was extracted from rat livers and mRNAconcentrations were determined by realtime detection RT-PCR analysisusing b-actin mRNA concentration for normalization. Values aremeans ± SD (n = 8). An asterisk (*) indicates a significant difference fromcontrol rats (P < 0.05).

706 S. Luci et al. / Biochemical and Biophysical Research Communications 350 (2006) 704–708

63

between cells treated with WY-14,643 and control cells(Fig. 2).

Discussion

In this study, rats and Fao cells were treated with thePPARa agonists clofibrate and WY-14,643, respectively.CPT-1 and CPT-2 have been identified as PPARa targetgenes [22–24], and the observation that expression of thesegenes was strongly up-regulated in the liver of rats and inFao cells by treatment with the agonists indicated PPARaactivation. The rat study confirms the previous observationthat clofibrate treatment increases the concentration of car-nitine in the liver of rats [9,10]. The novel finding of thisstudy is that clofibrate treatment caused a strong up-regu-lation of OCTN-2 (8.4-fold) and a less strong up-regulationof OCTN-1 (2.4-fold) whereas mRNA concentrations ofgenes encoding enzymes of hepatic carnitine biosynthesisremained unchanged. These observations strongly indicatethat the increased carnitine concentration in the liver ofrats treated with clofibrate was rather due to an increaseduptake of carnitine from the blood into the liver than toan increased synthesis of carnitine. This indication accordswith the observation that carnitine concentrations in plas-ma were reduced by clofibrate treatment, probably due toan increased uptake into cells. More than 95% of the totalcarnitine in the body is localized in the muscle which servesas a carnitine storage. When plasma carnitine concentra-tions are lowered such as by treatment with pivalate, carni-tine can be mobilized from the muscle in order to normalizeplasma carnitine concentrations [25]. The finding that theconcentration of acetyl carnitine, the storage form of carni-

tine, was reduced in gastrocnemius of rats treated withclofibrate indeed indicates that carnitine might have beenmobilized from muscle.

The in vitro study in which Fao cells were treated withWY-14,643 confirms most of the observations of the ratstudy. It is shown that treatment of cells with WY-14,643increased gene expression of OCTN-2 and increased theintracellular concentration of free carnitine whereas geneexpression of enzymes of hepatic carnitine biosynthesiswas not altered, too. The only disagreement between the celland the rat study was that expression of OCTN-1 was notup-regulated by the PPARa agonist in Fao cells. Thismay have two reasons: first, the effect of WY-14,643 inFao cells was generally weaker than the effect of clofibratein rats on the respective parameters; second, OCTN-1was generally less responsive to PPARa agonists thanOCTN-2. As gene expression of OCTN-1 remainedunchanged in Fao cells treated with WY-14,643, it can beconcluded that the increased concentration of free carnitinewas exclusively the result of an increased carnitine uptakeinto the cell by OCTN-2.

The cell culture study disproves the hypothesis that theincrease of the carnitine concentration in livers of ratstreated with clofibrate is caused by an increased hepaticcarnitine synthesis due to an increased availability ofTML derived from muscle [10]. Carnitine concentrationin Fao cells was increased by WY-14,643 although concen-trations of TML in the media of treated and control cellswere identical. This means that carnitine concentrationsin Fao cells were increased by WY-14,643 independent ofthe availability of TML from muscle or other tissues.

The observation that OCTN-2 expression was up-regu-lated and that carnitine concentration was increased in liv-er and cells treated with two different PPARa agonistsindicates that these effects were caused by PPARa activa-tion. This indication provides also an explanation for theobservation of increased hepatic carnitine concentrationsin fasted rats [15,16]. During fasting, non-esterified fattyacids are liberated from adipose tissue and act as activatorsof PPARa when they have entered the liver. Activation ofPPARa up-regulates many genes involved in hepatic mito-chondrial and peroxisomal b-oxidation of fatty acids tosupply acetyl-CoA used for the generation of ATP via cit-rate cycle and for the generation of ketone bodies, animportant fuel for the brain during fasting [17,26]. Thesemetabolic adaptations during fasting triggered by PPARaaim to minimize the use of protein and carbohydrates asfuel and allow mammals to survive long periods of energydeprivation. CPTs are rate-limiting for b-oxidation of fattyacids [22,24]. The up-regulation of CPTs, which is essentialfor the metabolic adaptations occurring during fasting,might increase the demand of carnitine in liver cells. Wepostulate that up-regulation of OCTNs by PPARa activa-tion is a means to supply liver cells with sufficient carnitinerequired for transport of excessive amounts of fatty acidsinto the mitochondrion, and therefore plays an importantrole in the adaptive response of liver metabolism to fasting.

Fig. 2. Effect of WY-14,643 treatment for either 6 or 20 h on the relativemRNA concentrations of carnitine palmitoyltransferases (CPT-1, CPT-2),organic cation transporters (OCTN-1, OCTN-2), and enzymes of carnitinebiosynthesis (trimethyllysine dioxygenase, TMLD; 4-N-trimethylamino-butyraldehyde dehydrogenase, TMABA-DH; c-butyrobetaine dioxygen-ase, BBD) in Fao cells. Fao cells were grown in culture medium untilsubconfluent state and were then incubated with 50 lM of the PPARaagonist WY-14,643 for 6 and 20 h, respectively. Control cells wereincubated with low-serum medium containing vehicle alone. Total RNAwas extracted from cells and mRNA concentrations were determined byrealtime detection RT-PCR analysis using b-actin mRNA concentrationfor normalization. mRNA concentrations of the genes in the treated cellsare shown relative to control cells (=1.00; dotted line) treated for 6 and20 h, respectively. An asterisk (*) indicates a significant difference fromcontrol cells (P < 0.05).

S. Luci et al. / Biochemical and Biophysical Research Communications 350 (2006) 704–708 707

64

In conclusion, the present study shows for the first timethat treatment of rats and rat Fao hepatoma cells withPPARa agonists clofibrate and WY-14,643, respectively,causes an up-regulation of OCTNs whereas expression ofgenes encoding enzymes involved in hepatic carnitine bio-synthesis remain unchanged. Up-regulation of OCTNsmay enhance carnitine uptake from blood or medium,respectively, into liver cells and this may be the reasonfor the increased carnitine concentrations in livers of ratstreated with clofibrate.

References

[1] J.D. McGarry, N.F. Brown, The mitochondrial carnitine palmitoyl-transferase system. From concept to molecular analysis, Eur. J.Biochem. 244 (1997) 1–14.

[2] E.P. Brass, Pivalate-generating prodrugs and carnitine homeostasis inman, Pharmacol. Rev. 54 (2002) 589–598.

[3] A. Steiber, J. Kerner, C.L. Hoppel, Carnitine: a nutritional,biosynthetic, and functional perspective, Mol. Aspects Med. 25(2004) 455–473.

[4] C.J. Rebouche, H. Seim, Carnitine metabolism and its regulation inmicroorganisms and mammals, Annu. Rev. Nutr. 18 (1998) 39–61.

[5] C.L. Hoppel, A.T. Davis, Inter-tissue relationships in the synthesisand distribution of carnitine, Biochem. Soc. Trans. 14 (1986) 673–674.

[6] F.M. Vaz, R.J. Wanders, Carnitine biosynthesis in mammals,Biochem. J. 361 (2002) 417–429.

[7] A.M. Evans, G. Fornasini, Pharmacokinetics of L-carnitine, Clin.Pharmacokinet. 42 (2003) 941–967.

[8] I. Tein, Carnitine transport: pathophysiology and metabolism ofknown defects, J. Inherit. Metab. Dis. 26 (2003) 147–169.

[9] H.S. Paul, S.A. Adibi, Paradoxical effects of clofibrate on liver andmuscle metabolism in rats. Induction of myotonia and alteration offatty acid and glucose oxidation, J. Clin. Invest. 64 (1979) 405–412.

[10] H.S. Paul, C.E. Gleditsch, S.A. Adibi, Mechanism of increasedhepatic concentration of carnitine by clofibrate, Am. J. Physiol. 251(1986) E311–E315.

[11] K. Schoonjans, J. Peinado-Onsurbe, A.M. Lefebvre, R.A. Heyman,M. Briggs, S. Deeb, B. Staels, J. Auwerx, PPARa and PPARcactivators direct a distinct tissue-specific transcriptional response via aPPRE in the lipoprotein lipase gene, EMBO J. 15 (1996) 5336–5348.

[12] J.A. Kramer, E.A. Blomme, R.T. Bunch, J.C. Davila, C.J. Jackson,P.F. Jones, K.L. Kolaja, S.W. Curtiss, Transcription profilingdistinguishes dose-dependent effects in the livers of rats treated withclofibrate, Toxicol. Pathol. 31 (2003) 417–431.

[13] S. Mandard, M. Muller, S. Kersten, Peroxisome proliferator receptora target genes, Cell Mol. Life Sci. 61 (2004) 393–416.

[14] A.L. Slitt, N.J. Cherrington, D.P. Hartley, M.T. Leazer, C.D.Klaassen, Tissue distribution and renal developmental changes inrat organic cation transporter mRNA levels, Drug Metab. Dispos. 30(2002) 212–219.

[15] J.D. McGarry, C. Robles-Valdes, D.W. Foster, Role of carnitine inhepatic ketogenesis, Proc. Natl. Acad. Sci. USA 72 (1975) 4385–4388.

[16] E.P. Brass, C.L. Hoppel, Carnitine metabolism in the fasting rat, J.Biol. Chem. 253 (1978) 2688–2693.

[17] S. Kersten, J. Seydoux, J.M. Peters, F.J. Gonzalez, B. Desvergne,W. Wahli, Peroxisome proliferator-activated receptor a mediatesthe adaptive response to fasting, J. Clin. Invest. 103 (1999) 1489–1498.

[18] T. Mossman, Rapid colorimetric assay for cellular growth andsurvival: application to proliferation and cytotoxicity assays, J.Immunol. Methods 65 (1983) 55–63.

[19] L. Vernez, M. Wenk, S. Krahenbuhl, Determination of carnitine andacylcarnitines in plasma by high-performance liquid chromatogra-phy/electrospray ionization ion trap tandem mass spectrometry,Rapid Commun. Mass Spectrom. 18 (2004) 1233–1238.

[20] B. Konig, K. Eder, Differential action of 13-HPODE on PPARadownstream genes in rat Fao and human HepG2 hepatoma cell lines,J. Nutr. Biochem. 17 (2006) 410–418.

[21] M.W. Pfaffl, A new mathematical model for relative quantification inreal-time RT-PCR, Nucleic Acids Res. 29 (2001) e45.

[22] J.M. Brandt, F. Djouadi, D. Kelly, Fatty acids activate transcriptionof the muscle carnitine palmitoyltransferase I gene in cardiacmyocytes via the peroxisome proliferator-activated receptor alpha,J. Biol. Chem. 273 (1998) 23786–23792.

[23] T. Hashimoto, T. Fujita, N. Usuda, W. Cook, C. Qi, J.M. Peters, F.J.Gonzalez, A.V. Yeldandi, M.S. Rao, J.K. Reddy, Peroxisomal andmitochondrial fatty acid beta-oxidation in mice nullizygous for bothperoxisome proliferator-activated receptor alpha and peroxisomalfatty acyl-CoA oxidase. Genotype correlation with fatty liver pheno-type, J. Biol. Chem. 274 (1999) 19228–19236.

[24] C. Mascaro, E. Acosta, J.A. Ortiz, P.F. Marrero, F.G. Hegardt, D.Haro, Control of human muscle-type carnitine palmitoyltransferase Igene transcription by peroxisome proliferator-activated receptor, J.Biol. Chem. 273 (1998) 8560–8563.

[25] H. Nakajima, N. Kodo, F. Inoue, Z. Kizaki, S. Nukina, A. Kinugasa,T. Sawada, Pivalate affects carnitine status but causes no severemetabolic changes in rat liver, J. Nutr. 126 (1996) 1683–1687.

[26] Y. Cheon, T.Y. Nara, M.R. Band, J.E. Beever, M.A. Wallig, M.T.Nakamura, Induction of overlapping genes by fasting and a perox-isome proliferator in pigs: evidence of functional PPARalpha innonproliferating species, Am. J. Physiol. Integr. Comp. Physiol. 288(2005) R1525–R1535.

708 S. Luci et al. / Biochemical and Biophysical Research Communications 350 (2006) 704–708

65

Dietary oxidised fat up regulates the expression of organic cation transporters

in liver and small intestine and alters carnitine concentrations in liver, muscle

and plasma of rats

Alexander Koch, Bettina Konig, Sebastian Luci, Gabriele I. Stangl and Klaus Eder*

Institute of Agricultural and Nutritional Sciences, Martin-Luther-University of Halle-Wittenberg, Emil-Abderhalden-Strasse 26,

D-06108 Halle (Saale), Germany

(Received 19 December 2006 – Revised 3 April 2007 – Accepted 12 April 2007)

It has been shown that treatment of rats with clofibrate, a synthetic agonist of PPARa, increases mRNA concentration of organic cation transpor-

ters (OCTN)-1 and -2 and concentration of carnitine in the liver. Since oxidised fats have been demonstrated in rats to activate hepatic PPARa, we

tested the hypothesis that they also up regulate OCTN. Eighteen rats were orally administered either sunflower-seed oil (control group) or an oxi-

dised fat prepared by heating sunflower-seed oil, for 6 d. Rats administered the oxidised fat had higher mRNA concentrations of typical PPARa

target genes such as acyl-CoA oxidase, cytochrome P450 4A1 and carnitine palmitoyltransferases-1A and -2 in liver and small intestine than con-

trol rats (P,0·05). Furthermore, rats treated with oxidised fat had higher hepatic mRNA concentrations of OCTN1 (1·5-fold) and OCTN2 (3·1-

fold), a higher carnitine concentration in the liver and lower carnitine concentrations in plasma, gastrocnemius and heart muscle than control rats

(P,0·05). Moreover, rats administered oxidised fat had a higher mRNA concentration of OCTN2 in small intestine (2·4-fold; P,0·05) than con-

trol rats. In conclusion, the present study shows that an oxidised fat causes an up regulation of OCTN in the liver and small intestine. An increased

hepatic carnitine concentration in rats treated with the oxidised fat is probably at least in part due to an increased uptake of carnitine into the liver

which in turn leads to reduced plasma and muscle carnitine concentrations. The present study supports the hypothesis that nutrients acting as

PPARa agonists influence whole-body carnitine homeostasis.

Carnitine: Oxidised fat: Peroxisome proliferator-activated receptor-a: Organic cation transporters

Carnitine (L-3-hydroxy-4-N-N-N-trimethylaminobutyrate) isan essential metabolite that has a number of indispensablefunctions in intermediary metabolism. The most prominentfunction lies in its role in the transport of activated long-chain fatty acids from the cytosol to the mitochondrialmatrix where b-oxidation takes place1 – 3. All tissues that usefatty acids as a fuel source require carnitine for normal func-tion. Carnitine is derived from dietary sources and endogenousbiosynthesis4. Carnitine biosynthesis involves a complexseries of reactions involving several tissues5. Lysine providesthe carbon backbone of carnitine. Lysine in protein peptidelinkages undergoes methylation of the e-amino group toyield trimethyllysine, which is released upon protein degra-dation. Muscle is the major source of trimethyllysine. Thereleased trimethyllysine is further oxidised to butyrobetaineby the action of trimethyllysine dioxygenase, 3-hydroxy-N-tri-methyllysine aldolase and 4-N-trimethylaminobutyraldehydedehydrogenase. Butyrobetaine is hydroxylated by g-butyro-betaine dioxygenase to form carnitine. The last reactionwhich is rate-limiting for carnitine synthesis occurs primarilyin the liver and kidneys6 (see Fig. 1).

Distribution of carnitine within the body and intracellularhomeostasis of carnitine are controlled by organic cation

transporters (OCTN) which belong to the solute carrier (SLC)22A family, localised to the apical membrane of cells7,8.Three OCTN have been identified so far: OCTN1, OCTN2and OCTN39 – 11. OCTN are polyspecific; they transport severalcations and L-carnitine12,13. Carnitine transport by OCTN1 andOCTN2 is Na dependent whereas that by OCTN3 is Na indepen-dent11. OCTN1 and OCTN2 are expressed in several tissuessuch as kidney, intestine, skeletal muscle, heart, liver andbrain11,14,15. In contrast, OCTN3 is expressed exclusively inthe testes and kidneys11. Among the three OCTN, OCTN3 hasthe highest specificity for carnitine; OCTN1 has the lowestone11. OCTN operate on the intestinal absorption and renal reab-sorption of carnitine and play a major role in tissue distributionby catalysing the uptake of carnitine into body cells. Due to itshigh binding affinity for carnitine and its wide expression,OCTN2 seems to be the most physiologically important carni-tine transporter. OCTN1 contributes less to carnitine transportthan OCTN2 due to its low carnitine transport activity.OCTN3 may be important for carnitine uptake into the testes,and may contribute to the reabsorption of carnitine in the kid-neys11. The fact that inborn or acquired defects of OCTN leadto primary or secondary systemic carnitine deficiency demon-strates their essential role in carnitine homeostasis8.

*Corresponding author: Professor Dr Klaus Eder, fax þ345 5527124, email [email protected]

Abbreviations: CPT, carnitine palmitoyltransferase; Cyp, cytochrome P450; OCTN, organic cation transporter.

British Journal of Nutrition (2007), 98, 882–889 doi: 10.1017/S000711450775691Xq The Authors 2007

British

Journal

ofNutrition

66

It has been shown previously that starvation or treatment ofrats with clofibrate increases the concentration of carnitine inthe liver16 – 18. Both starvation and clofibrate treatment lead toan activation of PPARa, a transcription factor belonging to thenuclear hormone receptor superfamily19. We have recentlyshown that activation of PPARa by clofibrate treatmentcauses an up regulation of OCTN1 and OCTN2 in ratliver20. These results strongly indicated that increased carni-tine concentrations in livers of rats starved or treated with clo-fibrate were due to increased uptake of carnitine from bloodinto the liver. Indeed, plasma carnitine concentrations werereduced in rats treated with clofibrate which may be causedby an increased uptake into the liver20.

In addition to synthetic agonists, several naturally occurringcompounds are able to activate PPARa in vivo. Recently, weand others have shown that dietary oxidised fats prepared bythe heating of vegetable oils activate hepatic PPARa in ratsand pigs21 – 25. In the present study, we tested the hypothesis

that oxidised fats are also able to up regulate the expressionof OCTN (OCTN1, OCTN2) in the liver due to their abilityto activate PPARa and thereby increase hepatic carnitineconcentration. For this end, we performed an experimentwith growing rats as an animal model, according to a previousstudy dealing with the effects of an oxidised oil on PPARaactivation24.

More than 95 % of the total carnitine in the body is localisedin the muscle which serves as a carnitine storage2. Whenplasma carnitine concentrations are lowered, such as by treat-ment with pivalate, carnitine is mobilised from the muscle inorder to normalise plasma carnitine concentrations26. There-fore, an increased uptake of carnitine from the blood intothe liver by up regulation of hepatic OCTN should lead to amobilisation of carnitine storage in the muscle. To investigatethis, we also determined carnitine concentrations in skeletalmuscle and heart of the rats.

OCTN1 and OCTN2 are also highly expressed in the intes-tine and particularly OCTN2 plays an important role in theabsorption of L-carnitine from the diet15,27,28. As the smallintestine also has a high expression of PPARa, it seems poss-ible that an oxidised fat could increase the gene expression ofOCTN also in the small intestine via an activation of PPARa.Besides OCTN, the amino acid transporter ATB0þ is involvedin the intestinal absorption of carnitine from the diet27,29.In order to obtain information whether PPARa activation bysynthetic or native agonists could influence intestinal carnitineabsorption, we also determined mRNA concentration ofATB0þ in small intestine.


Animal experiment

Male Sprague–Dawley rats, aged 5 weeks old, supplied byCharles River (Sulzfeld, Germany) with an average initialbody weight of 115 (SD 14) g were randomly assigned totwo groups of nine rats each. They were kept individually inMacrolon cages in a room controlled for temperature(22 ^ 28C), relative humidity (50–60 %) and light (12 hlight–dark cycle). All experimental procedures describedfollowed established guidelines for the care and handling oflaboratory animals and were approved by the council ofSaxony-Anhalt. The animals received either 2 ml fresh sun-flower-seed oil (control group) or oxidised sunflower-seedoil (see Preparation of the oxidised fat) by oral administrationonce per d 2 h after the beginning of the light cycle. After-wards, they obtained their daily food ration. All rats werefed a commercial standard basal diet (Altromin 1324; Altro-min GmbH, Lage, Germany). Concentration of total carnitinein the basal diet was 22mmol/kg. To standardise food intake,diet intake was controlled. Each rat in the experiment received12 g diet/d. This amount of diet which is approximately 20 %below the amount of diet rats would consume ad libitum wascompletely ingested by all rats. Thus, the diet intake was iden-tical in all the rats within this experiment. Water was availablead libitum from nipple drinkers during the whole experiment.At day 6 of treatment, rats received the last dose of fresh oroxidised fat and 9 g diet and were killed 4 h later by decapi-tation under light anaesthesia with diethyl ether. Blood wascollected into heparinised polyethylene tubes. Liver, heart

Fig. 1. Schematic diagram of carnitine biosynthesis from trimethyllysine

(TML) (according to Vaz & Wanders6). TML is oxidised to butyrobetaine

by trimethyllysine dioxygenase (TMLD), 3-hydroxy-N-trimethyllysine

aldolase (HTMLA) and 4-N-trimethylaminobutyraldehyde dehydrogenase

(TMABA-DH). In the last rate-limiting step, butyrobetaine is hydroxylated to

L-carnitine by g-butyrobetaine dioxygenase (BBD). HTML, 3-hydroxy-N-tri-

methyllysine; TMABA, 4-N-trimethylaminobutyraldehyde.

Oxidised fat and organic cation transporters 883

British

Journal

ofNutrition

67

and gastrocnemius muscles were quickly removed, frozenwith liquid N2 and stored at 2808C pending further analysis.Plasma was obtained by centrifugation of the blood (1100 g;10 min; 48C) and stored at 2208C. Liver samples for RNAisolation and lipid extraction were snap-frozen in liquid N2

and stored at 2808C. The small intestine was rapidly excised,washed with cold 0·9 % NaCl (w/v) and mucosal scrapingswere obtained from the jejunum (defined by length), snap-frozen and stored at 2808C for RNA extraction.

Preparation of the oxidised fat

The thermoxidised oil was prepared by heating sunflower-seedoil obtained from a local supermarket in an electric fryer (SaroGastro-Products GmbH, Emmerich, Germany) for 25 d at608C. Throughout the heating process, air was continuouslybubbled into the fat. The extent of lipid peroxidation was deter-mined by assaying the peroxide value30, concentration of thio-barbituric acid-reactive substances31 and conjugated dienes32,acid values30, the percentage of total polar compounds33 andthe concentration of total carbonyls34. The oxidised fat hadmuch higher concentrations of peroxides (126-fold), conjugateddienes (.2740-fold), thiobarbituric acid-reactive substances(12-fold), total carbonyls (33-fold), polar compounds (5-fold)and a higher acid value (15-fold) than the fresh fat (Table 1).

Carnitine analysis

Carnitine was determined as [3H]acetyl-carnitine after the ester-ification of non-esterified carnitine by carnitine acyltransferaseaccording to McGarry & Foster35 with modifications proposedby Parvin & Pande36 and Christiansen & Bremer37. Plasmasamples were used directly for the determination of the total car-nitine after alkaline hydrolysation as described for the tissuesamples below. Tissue samples were freeze dried and milled.Then 100 mg liver or 50 mg muscle powder were sonificatedin 5 ml water for 15 min. Samples were centrifuged (12 000 g;5 min) and non-esterified carnitine in the supernatant fractionwas measured. For the determination of the total carnitine thesamples were hydrolysed before the centrifugation. For this,10 ml 0·2 M-potassium hydroxide were added, the sampleswere incubated at 308C for 1 h and then neutralised by theaddition of 0·2 M-HCl. Carnitine esterification was done in afinal volume of 1 ml containing 0·1 M-HEPES (pH 7·4), 2 mM-N-ethylmaleimide, 1·25 mM-EDTA, 25mM-[3H]acetyl-CoA(29·4 MBq/mmol; GE Healthcare, Buckinghamshire, UK) and1 U carnitine acyltransferase (Roche Diagnostic, Mannheim,Germany) for 30 min at room temperature. [3H]acetyl-CoA not

consumed by the reaction was bound to Dowex 1-X 8 and separ-ated by centrifugation. Carnitine concentration was calculatedusing the radioactivity of the supernatant fraction measured ina liquid scintillation counter and corrected for non-specificradioactivity.

Reverse transcriptase polymerase chain reaction analysis

Total RNA was isolated from rat livers and mucosa scrapings,respectively, by TRIZOLe reagent (Sigma-Aldrich, Steinheim,Germany) according to the manufacturer’s protocol. cDNA syn-thesis was carried out as described38. The mRNA concentrationof genes was measured by real-time detection PCR usingSYBRw Green I and the Rotor Gene 2000 system (CorbettResearch, Mortlake, Australia). Real-time detection PCR wasperformed with 1·25 U Taq DNA polymerase (Promega, Man-nheim, Germany), 500mM-dNTP and 26·7 pmol of the specificprimers (Operon Biotechnologies, Cologne, Germany; Table2). Annealing temperature for all primers was 608C. For determi-nation of mRNA concentration a threshold cycle (Ct) and ampli-fication efficiency was obtained from each amplification curveusing the software RotorGene 4·6 (Corbett Research). Calcu-lation of the relative mRNA concentration was made using theDDCt method as previously described39. The housekeepinggene glyceraldehyde-3-phosphate dehydrogenase was used fornormalisation. mRNA concentration of glyceraldehyde-3-phos-phate dehydrogenase was not influenced by the treatment of ratswith oxidised fat.

Statistical analysis

Means of the treatment and control groups were compared byan unpaired t test using the Minitab Statistical Software (Mini-tab, State College, PA, USA). Differences with P,0·05 wereconsidered to be significant.

Results

Final weights and body-weight gains of the rats

Final body weights of rats treated with the oxidised fat (133(SD 14) g) were not significantly different from the controlrats (144 (SD 14) g) (nine rats for each group). However,rats treated with the oxidised fat had a lower body-weightgain (17·5 (SD 6·4) g) over the feeding period than the controlrats (29·7 (SD 4·5) g) (nine rats for each group; P,0·05).

mRNA concentrations of acyl-CoA oxidase, cytochrome P450-4A1, carnitine palmitoyltransferases-1A and -2, organiccation transporters-1 and -2 and enzymes involved in hepaticcarnitine synthesis (trimethyllysine dioxygenase, 4-N-trimethylaminobutyraldehyde dehydrogenase andg-butyrobetaine dioxygenase) in the liver

Rats treated with the oxidised fat had higher mRNA concen-trations of acyl-CoA oxidase, cytochrome P450 (Cyp)-4A1,carnitine palmitoyltransferase (CPT)-2, OCTN1 and OCTN2in the liver than control rats (P,0·05); mRNA concentrationof CPT1A, however, was not different in the rats treatedwith oxidised fat from the control rats (Fig. 2). Rats treatedwith the oxidised fat had a higher mRNA concentration oftrimethyllysine dioxygenase in the liver than control

Table 1. Concentrations of various lipid oxidation productsin the fats*

Oxidation product Fresh fat Oxidised fat

Peroxide value (mEq O2/kg) 3·0 378·6Conjugated dienes (mmol/kg) ,0·1 273·6TBARS (mmol/kg) 1·1 13·1Total carbonyls (mmol/kg) 2·9 96·9Total polar compounds (%) 5·1 27·8Acid value (g KOH/kg) 0·4 5·8

TBARS, thiobarbituric acid-reactive substances.* Data are the results of single measurements.

A. Koch et al.884

British

Journal

ofNutrition

68

rats (P,0·05; Fig. 2). mRNA concentrations of 4-N-trimethy-laminobutyraldehyde dehydrogenase and g-butyrobetainedioxygenase in the liver, however, did not differ betweenthe two groups of rats (Fig. 2).

mRNA concentrations of acyl-CoA oxidase, cytochrome P450-4A1, carnitine palmitoyltransferases-1A and -2, organiccation transporters-1 and -2 and ATB0þ in the small intestine

Rats treated with oxidised fat had higher mRNA concentrationsof acyl-CoA oxidase, Cyp4A1, CPT1A, CPT2 and OCTN2(P,0·05), and they tended to have a higher mRNA concen-tration of OCTN1 (P¼0·066) in the small intestine comparedwith control rats (Fig. 3). mRNA concentration of ATB0þ inthe small intestine was reduced in the rats fed the oxidised fatcompared with those fed the fresh fat (P,0·05; Fig. 3).

Carnitine concentrations in liver, plasma and muscle

Rats treated with the oxidised fat had a higher carnitine con-centration in the liver than control rats (P,0·05, Fig. 4).

Plasma carnitine concentration was lower in the rats treatedwith oxidised fat (18·8 (SD 3·1) mmol/l) than in the controlgroup (28·4 (SD 4·3) mmol/l) (nine rats for each group;P,0·05). Rats treated with oxidised fat also had lower carni-tine concentrations in gastrocnemius and heart muscle thancontrol rats (P,0·05; Fig. 4).

Discussion

We have recently found that treatment with clofibrate causes astrong up regulation of OCTN2, and a less strong up regu-lation of OCTN1, in the liver of rats which was accompaniedby an increased hepatic carnitine concentration20. This effectwas probably caused by PPARa activation. In the presentstudy, we investigated the hypothesis that oxidised fats areable to exert similar effects due to their ability to activatePPARa. Hydroxy- and hydroperoxy fatty acids such as hy-droxyoctadecadienoic and hydroperoxyoctadecadienoic acidoccurring in oxidised fats are very potent PPARa ago-nists38,40,41. These fatty acids are produced during the earlystage of lipid peroxidation. Since they are unstable anddecompose at high temperatures, fats treated at low tempera-ture have much higher concentrations of these primary lipid

Table 2. Characteristics of the primers used in reverse transcriptase polymerase chain reaction analysis

Gene Forward primer (from 50 to 30) Reverse primer (from 50 to 30) bp NCBI GenBank

ACO CTTTCTTGCTTGCCTTCCTTCTCC GCCGTTTCACCGCCTCGTA 415 NM_017340ATB0þ ATCCGGAAGCACTAGCTCAA CCCAGTAAATTCCAGCCTGA 237 NM_001037544BBD ATTCTGCAAAAGCTCGGAAA CTCCTTGGAGTCCTGCTCTG 183 NM_022629Cyp4A1 CAGAATGGAGAATGGGGACAGC TGAGAAGGGCAGGAATGAGTGG 460 NM_175837CPT1A GGAGACAGACACCATCCAACATA AGGTGATGGACTTGTCAAACC 416 NM_031559CPT2 TCCTCGATCAAGATGGGAAC GATCCTTCATCGGGAAGTCA 237 NM_012930GAPDH GCATGGCCTTCCGTGTTCC GGGTGGTCCAGGGTTTCTTACTC 337 BC059110OCTN1 AGCATTTGTCCTGGGAACAG ACTCAGGGATGAACCACCAG 200 NM_022270OCTN2 CCTCTCTGGCCTGATTGAAG CTCCGCTGTGAAGACGTACA 226 NM_012930TMLD GCCCTGTGGCATTCAAGTAT GGTCCAACCCCTATCATGTG 201 AF374406TMABA-DH TTTGAGACTGAAGCCGAGGT CACCGGGCTGACGTTATAGT 156 NM_022273

ACO, acyl-CoA oxidase; BBD, g-butyrobetaine dioxygenase; Cyp, cytochrome P450; CPT, carnitine palmitoyltransferase; GAPDH, glyceralde-hyde-3-phosphate dehydrogenase; OCTN, organic cation transporter; TMLD, trimethyllysine dioxygenase; TMABA-DH, 4-N-trimethylaminobu-tyraldehyde dehydrogenase.

Fig. 2. Effect of an oxidised fat on the relative mRNA concentrations of acyl-CoA

oxidase (ACO), cytochrome P450 (Cyp)-4A1, carnitine palmitoyltransferases

(CPT)-1A and -2, organic cation transporters (OCTN)-1 and -2,

trimethyllysine dioxygenase (TMLD), 4-N-trimethylaminobutyraldehyde dehydro-

genase (TMABA-DH) and g-butyrobetaine dioxygenase (BBD) in the liver of

rats. Rats were treated orally with 2 ml oxidised fat (A) or fresh fat (B;

control ¼ 1·00) for 6 d. Total RNA was extracted from rat livers and mRNA

concentrations were determined by real-time detection RT-PCR analysis using

glyceraldehyde-3-phosphate dehydrogenase mRNA concentration for normali-

sation. Values are means, with standard deviations represented by vertical bars

(n 9). *Mean value was significantly different from that of the control rats

(P,0·05).

Fig. 3. Effect of an oxidised fat on the relative mRNA concentrations of acyl-

CoA oxidase (ACO), cytochrome P450 (Cyp)-4A1, carnitine palmitoyltrans-

ferases (CPT)-1A and -2, organic cation transporters (OCTN)-1 and -2 and

amino acid transporter ATB0þ in the small intestine of rats. Rats were treated

orally with 2 ml oxidised fat (A) or fresh fat (B; control ¼ 1·00) for 6 d. Total

RNA was extracted from mucosal scrapings and mRNA concentrations were

determined by real-time detection RT-PCR analysis using glyceraldehyde-3-

phosphate dehydrogenase mRNA concentration for normalisation. Values

are means, with standard deviations represented by vertical bars (n 9).

*Mean value was significantly different from that of the control rats (P,0·05).


British

Journal

ofNutrition

69

peroxidation products than fats treated at high temperature24.This is the reason why we used a fat treated at a relativelylow temperature for a long period. The high peroxide valueand the high concentration of conjugated dienes indicate thatthis fat indeed had high concentrations of hydroxy- and hydro-peroxy fatty acids which may be particularly responsible forthe PPARa-activating effects of oxidised fats. To ensurethat all rats obtained the same dose of oxidised fat, it wasadministered orally. The oxidised and fresh fat, respectively,accounted for about 25 % of total energy of the total dailyfeed. Since it was observed in a previous rat study that evenshort-term application of a PPARa agonist led to the typicalchanges known for PPARa activation such as up regulationof classical target genes involved in b-oxidation and reductionof TAG concentration38, we decided to give the oxidised fatover a relatively short time of 6 d. It has been shown thatthe intake of oxidised fats could cause a reduction of thefood intake in rats which could cause secondary effectswhich interact with the effect of treatment42,43. To ensure anidentical food intake in both groups of rats, we used acontrolled feeding system in which each rat consumed 12 gdiet /d. This amount of diet is slightly below that that ratswould consume ad libitum but in clear excess of that necess-ary to meet the maintenance energy requirement (which isapproximately 6 g/d44) and ensures an adequate growth ofthe rats. It is known that fasting causes an activation ofPPARa due to the release of NEFA from the adiposetissue45. To avoid PPARa activation due to an insufficientsupply of energy, all the rats received their last portion ofdiet 4 h before decapitation. Therefore, we can exclude thepossibility that PPARa was also activated in the controlgroup fed the fresh fat. The finding that rats fed the oxidisedfat gained less weight during the experimental period thanthose fed the fresh fat although both groups received an iden-tical amount of diet indicates that the oxidised fat impaired thefeed conversion ratio. This finding agrees with other reportswhich also showed that feeding of oxidised fats impairs thegrowth of rats46 – 49. We did not investigate the reason forthis. Previous studies, however, have shown that oxidisedfats lower the digestibility of nutrients46,50 and this may bethe reason for the reduced body-weight gains of the rats fedthe oxidised fat observed in the present study. However, as

rats fed the oxidised fat appeared quite normal, we assumethat the oxidised fat did not cause general toxicity.

The finding of increased mRNA concentrations of the typi-cal PPARa downstream genes acyl-CoA oxidase, Cyp4A1,CPT1A and CPT2 (for a review, see Mandard et al.45) inliver and intestine indeed indicates that the oxidised fatcaused an activation of PPARa in both liver and intestine ofthe rats. This indication agrees with recent studies in ratsand pigs which also showed that intake of oxidised fatsleads to an activation of PPARa in the liver21 – 25.

The present study shows further that treatment of rats withan oxidised fat caused the same alterations as observed forclofibrate20, namely increased hepatic mRNA concentrationsof OCTN1 and OCTN2 and an increased hepatic carnitineconcentration. Considering that a similar up regulation ofOCTN1 and OCTN2 was observed in the liver of rats treatedwith the synthetic PPARa agonist clofibrate and in rat hepa-toma cells treated with the more potent and selectivePPARa agonist WY 14,64320, we propose that the oxidisedfat up regulated OCTN in the liver also by PPARa activation.

In rat liver, OCTN1 and OCTN2 are highly expressed15.Both of them are able to transport carnitine into the livercell51,52. However, it has been shown that OCTN2 has ahigher carnitine transport activity than OCTN111. For thatreason and as mRNA concentration of OCTN2 was morestrongly increased by the oxidised fat than that of OCTN1,we assume that increased hepatic carnitine concentrations inrats treated with oxidised fat were caused mainly by anincreased uptake of carnitine via OCTN2. Plasma carnitineconcentrations are regulated by several events, namely intesti-nal absorption from the diet, renal excretion, endogenous syn-thesis in the liver and kidneys and movement of carnitinebetween plasma and tissues53. We have not studied the phar-macokinetics of carnitine but it seems plausible that reducedplasma concentrations of carnitine in rats fed the oxidisedfat may at least in part be due to an enhanced uptake intothe liver. We measured mRNA concentrations of OCTNonly in liver and small intestine; however, it is possible thatthey were increased also in other tissues in rats fed the oxi-dised fat. Therefore, an increased uptake of carnitine intoother tissues besides liver could also contribute to the reducedplasma carnitine concentrations. In the kidney, OCTN2 func-tions to reabsorb carnitine from the urine13,54. An up regu-lation of OCTN2 in kidney would be expected to reduceurinary excretion of carnitine which in turn results in anincreased plasma carnitine concentration. However, theeffect of oxidised fats on the gene expression of OCTN inthose tissues and their consequences on whole-body carnitinehomeostasis should be determined in future studies.

In the present study we also determined mRNA concen-trations of various enzymes involved in hepatic carnitinebiosynthesis in the liver which belongs like the kidney tothe tissues being able to synthesise carnitine6. It was foundthat oxidised fat treatment led to a moderate up regulationof trimethyllysine dioxygenase while mRNA concentrationsof 4-N-trimethylaminobutyraldehyde dehydrogenase andg-butyrobetaine dioxygenase, the rate-limiting enzyme of car-nitine biosynthesis6, remained unchanged by the treatment.This finding shows that PPARa activation by the oxidisedoil does not up regulate the gene expression of enzymesinvolved in hepatic carnitine synthesis. Nevertheless, it is

Fig. 4. Effect of an oxidised fat on the concentrations of total carnitine in

liver, gastrocnemius and heart. Rats were treated orally with 2 ml oxidised fat

(A) or fresh fat (B; control) for 6 d. Values are means, with standard devi-

ations represented by vertical bars (n 9). *Mean value was significantly differ-

ent from that of the control rats (P,0·05).

A. Koch et al.886

British

Journal

ofNutrition

70

possible that carnitine hepatic biosynthesis was increased inrats treated with oxidised fat. The liver has a high capacityto convert g-butyrobetaine into carnitine6. As OCTN2 has ahigh affinity for g-butyrobetaine10,11 it is likely that anincreased expression of OCTN2 may have led to an increaseduptake of g-butyrobetaine from plasma into the liver which interm may have stimulated synthesis of carnitine in the liver.This assumption, however, has to be proven in further studies.

Muscle contains more than 95 % of whole-body carnitineand serves as a carnitine storage2. When plasma carnitine con-centrations are lowered, such as by treatment with pivalate,carnitine can be mobilised from the muscle in order to normal-ise plasma carnitine concentrations26. Therefore, we expectedthat a reduced plasma carnitine concentration may lead to areduction of the carnitine concentration in muscle. The findingthat the concentration of carnitine was reduced in gastrocne-mius and heart muscle of rats treated with oxidised fatindeed suggests that carnitine might have been mobilisedfrom muscle. In rats treated with clofibrate, a reduction ofmuscle carnitine concentration has also been found20. Areduced carnitine concentration in muscle could also be dueto a reduced uptake of carnitine due to a decreased activityof OCTN, which, however, is unlikely with respect to the find-ing that OCTN in liver were up regulated in rats fed the oxi-dised fat. As muscle also has a high expression of PPARa, weexpect that the expression of OCTN in muscle was increasedrather than reduced by the dietary oxidised fat.

The present study further shows that a dietary oxidised fatleads to an up regulation of OCTN2 in the small intestine.As PPARa target genes (acyl-CoA oxidase, CYP4A1,CPT1a, CPT2) in the intestine were also up regulated in ratsfed the oxidised fat, we assume that the increased expressionof OCTN in intestine was also caused by activation ofPPARa. As intestinal OCTN localised in the apical membraneof mucosa cells are able to transport carnitine from the dietinto the cell27,28, an increased expression of these transportersmay enhance their capacity to absorb carnitine. However, asATB0þ, another transporter involved in the intestinal absorp-tion of carnitine27, was down regulated in rats fed the oxidisedfat, it is difficult to draw conclusions about the whole intesti-nal absorption of carnitine from the diet. Nevertheless, theobserved up regulation of intestinal OCTN may be relevantbecause they are polyspecific and do not only transport carni-tine from the intestinal lumen into the mucosa cell but are alsoable to bind various drugs such as verapamil, spironolactoneor mildronate and other monovalent cations14,28,55 – 58. As oxi-dised fats increase the gene expression of OCTN in the smallintestine, it is possible that these fats also increase the absorp-tion of various drugs from the intestine.

The hypothesis that the up regulation of OCTN was causedby PPARa activation provides also an explanation for theobserved increased hepatic carnitine concentrations in fastedrats16,17. During fasting, NEFA are liberated from adiposetissue and act as activators of PPARa when they have enteredthe liver. Activation of PPARa up regulates many genesinvolved in hepatic mitochondrial and peroxisomal b-oxi-dation of fatty acids to supply acetyl-CoA used for the gener-ation of ATP via the citrate cycle and for the generation ofketone bodies, an important fuel for the brain duringfasting59,60. These metabolic adaptations during fasting trig-gered by PPARa aim to minimise the use of protein and

carbohydrates as fuel and allow mammals to survive longperiods of energy deprivation. CPT are rate limiting for b-oxi-dation of fatty acids61,62. The up regulation of CPT, which isessential for the metabolic adaptations occurring during fast-ing, might increase the demand for carnitine in liver cells.We postulate that up regulation of OCTN by PPARa acti-vation is a means to supply liver cells with sufficient carnitinerequired for the transport of excessive amounts of fatty acidsinto the mitochondrion, and therefore plays an importantrole in the adaptive response of liver metabolism to fasting.

In conclusion, the present study shows that an oxidised fatcauses an up regulation of OCTN2 in the liver and small intes-tine of rats. As OCTN2 catalyses the uptake of carnitine intocells, these fats influence whole-body carnitine homeostasis.An increased hepatic carnitine concentration in rats treatedwith oxidised fat may be at least in part due to an increaseduptake of carnitine from blood into the liver. Since OCTN2binds not only carnitine but also various drugs, the possibilityexists that increased OCTN2 expression in the small intestinemay improve the absorption of various drugs.

References

1. McGarry JD & Brown NF (1997) The mitochondrial carnitine

palmitoyltransferase system. From concept to molecular anal-

ysis. Eur J Biochem 244, 1–14.

2. Brass EP (2002) Pivalate-generating prodrugs and carnitine

homeostasis in man. Pharmacol Rev 54, 589–598.

3. Steiber A, Kerner J & Hoppel CL (2004) Carnitine: a nutri-

tional, biosynthetic, and functional perspective. Mol Asp Med

25, 455–473.

4. Rebouche CJ & Seim H (1998) Carnitine metabolism and its

regulation in microorganisms and mammals. Annu Rev Nutr

18, 39–61.

5. Hoppel CL & Davis AT (1986) Inter-tissue relationships in the

synthesis and distribution of carnitine. Biochem Soc Trans 14,

673–674.

6. Vaz FM & Wanders RJ (2002) Carnitine biosynthesis in mam-

mals. Biochem J 361, 417–429.

7. Lahjouji K, Mitchell GA & Qureshi IA (2001) Carnitine trans-

port by organic cation transporters and systemic carnitine

deficiency. Mol Genet Metab 73, 287–297.

8. Tein I (2003) Carnitine transport: pathophysiology and metab-

olism of known defects. J Inherit Metab Dis 26, 147–169.

9. Tamai I, Yabuuchi H, Nezu J, Sai Y, Oku A, Shimane M &

Tsuji A (1997) Cloning and characterization of a novel human

pH-dependent organic cation transporter, OCTN1. FEBS Lett

419, 107–111.

10. Tamai I, Ohashi R, Nezu J, Yabuuchi H, Oku A, Shimane M,

Sai Y & Tsuji A (1998) Molecular and functional identification

of sodium ion-dependent, high affinity human carnitine trans-

porter OCTN2. J Biol Chem 273, 20378–20382.

11. Tamai I, Ohashi R, Nezu JI, Sai Y, Kobayashi D, Oku A,

Shimane M & Tsuji A (2000) Molecular and functional charac-

terization of organic cation/carnitine transporter family in mice.

J Biol Chem 275, 40064–40072.

12. Ohashi R, Tamai I, Yabuuchi H, Nezu JI, Oku A, Sai Y, Shimane

M & Tsuji A (1999) Naþ-dependent carnitine transport by organic

cation transporter (OCTN2): its pharmacological and toxicologi-

cal relevance. J Pharmacol Exp Ther 291, 778–784.

13. Ohashi R, Tamai I, Nezu Ji J, Nikaido H, Hashimoto N, Oku A,

Sai Y, Shimane M & Tsuji A (2001) Molecular and physiologi-

cal evidence for multifunctionality of carnitine/organic cation

transporter OCTN2. Mol Pharmacol 59, 358–366.


British

Journal

ofNutrition

71

14. Wu X, Huang W, Prasad PD, Seth P, Rajan DP, Leibach FH,

Chen J, Conway SJ & Ganapathy V (1999) Functional charac-

teristics and tissue distribution pattern of organic cation trans-

porter 2 (OCTN2), an organic cation/carnitine transporter.

J Pharmacol Exp Ther 290, 1482–1492.

15. Slitt AL, Cherrington NJ, Hartley DP, Leazer MT & Klaassen

CD (2002) Tissue distribution and renal developmental changes

in rat organic cation transporter mRNA levels. Drug Metab

Dispos 30, 212–219.

16. McGarry JD, Robles-Valdes C & Foster DW (1975) Role of

carnitine in hepatic ketogenesis. Proc Natl Acad Sci USA 72,

4385–4388.

17. Brass EP & Hoppel CL (1978) Carnitine metabolism in the fast-

ing rat. J Biol Chem 253, 2688–2693.

18. Paul HS & Adibi SA (1979) Paradoxical effects of clofibrate on

liver and muscle metabolism in rats. Induction of myotonia and

alteration of fatty acid and glucose oxidation. J Clin Invest 64,

405–412.

19. Schoonjans K, Peinado-Onsurbe J, Lefebvre AM, Heyman RA,

Briggs M, Deeb S, Staels B & Auwerx J (1996) PPARa and

PPARg activators direct a distinct tissue-specific transcriptional

response via a PPRE in the lipoprotein lipase gene. Embo J 15,

5336–5348.

20. Luci S, Geissler S, Konig B, Koch A, Stangl GI, Hirche F & Eder K

(2006) PPARa agonists up-regulate organic cation transporters in

rat liver cells. Biochem Biophys Res Commun 24, 704–708.

21. Chao P-M, Chao C-Y, Lin F-J & Huang C-J (2001) Oxidized

frying oil up-regulates hepatic acyl-CoA oxidase and cyto-

chrome P450 4A1 genes in rats and activates PPARa. J Nutr

131, 3166–3174.

22. Chao P-M, Hsu SC, Lin F-J, Li YJ & Huang C-J (2004) The

up-regulation of hepatic acyl-CoA oxidase and cytochrome

P450 4A1 mRNA expression by dietary oxidized frying oil

is comparable between male and female rats. Lipids 39,

233–238.

23. Chao PM, Yang MF, Tseng YN, Chang KM, Lu KS & Huang

CJ (2005) Peroxisome proliferation in liver of rats fed oxidized

frying oil. J Nutr Sci Vitaminol (Tokyo) 51, 361–368.

24. Sulzle A, Hirche F & Eder K (2004) Thermally oxidized dietary

fat upregulates the expression of target genes of PPAR a in rat

liver. J Nutr 134, 1375–1383.

25. Luci S, Konig B, Giemsa B, Huber S, Hause G, Kluge H, Stangl

GI & Eder K (2007) Feeding of a deep-fried fat causes PPARa

activation in the liver of pigs as a non-proliferating species. Br J

Nutr 97, 872–882.

26. Nakajima H, Kodo N, Inoue F, Kizaki Z, Nukina S, Kinugasa A &

Sawada T (1996) Pivalate affects carnitine status but causes no

severe metabolic changes in rat liver. J Nutr 126, 1683–1687.

27. Taylor PM (2001) Absorbing competition for carnitine. J

Physiol 532, 283.

28. Kato Y, Sugiura M, Sugiura T, Wakayama T, Kubo Y, Kobaya-

shi D, Sai Y, Tamai I, Iseki S & Tsuji A (2006) Organic cation/

carnitine transporter OCTN2 (Slc22a5) is responsible for

carnitine transport across apical membranes of small intestinal

epithelial cells in mouse. Mol Pharmacol 70, 829–837.

29. Nakanishi T, Hatanaka T, Huang W, Prasad PD, Leibach FH,

Ganapathy ME & Ganapathy V (2001) Naþ- and Cl2-coupled

active transport of carnitine by the amino acid transporter

ATB0þ from mouse colon expressed in HRPE cells and Xeno-

pus oocytes. J Physiol 532, 297–304.

30. Deutsche Gesellschaft fur Fettwissenschaften (1994) Einheits-

methoden zur Untersuchung von Fetten, Fettprodukten, Tensiden

und verwandten Stoffen. Stuttgart, Germany: Wissenschaftliche

Verlagsgesellschaft.

31. Sidwell CG, Salwin H, Benca M & Mitchell JH Jr (1954) The

use of thiobarbituric acid as a measure of fat oxidation. J Am

Oil Chem Soc 31, 603–606.

32. Recknagel RO & Glende EA Jr (1984) Spectrophotometric

detection of lipid conjugated dienes. Methods Enzymol 105,

331–337.

33. International Union of Pure and Applied Chemistry (2000)

Determination of polar compounds, polymerized and oxidized

triacylglycerols, and diacylglycerols in oils and fats. Pure

Appl Chem 72, 1563–1575.

34. Endo Y, Li CM, Tagiri-Endo M & Fujimoto K (2001) A modi-

fied method for the estimation of total carbonyl compounds in

heated and frying oils using 2 propanol as a solvent. J Am Oil

Chem Soc 78, 1021–1024.

35. McGarry JD & Foster DW (1976) An improved and simplified

radioisotopic assay for the determination of free and esterified

carnitine. J Lipid Res 17, 277–281.

36. Parvin R & Pande SV (1977) Microdetermination of (-)carnitine

and carnitine acetyltransferase activity. Anal Biochem 79,

190–201.

37. Christiansen RZ & Bremer J (1978) Acetylation of tris(hydroxy-

methyl)aminomethane (Tris) and Tris derivatives by carnitine

acetyltransferase. FEBS Lett 86, 99–102.

38. Konig B & Eder K (2006) Differential action of 13-HPODE

on PPARa downstream genes in rat Fao and human HepG2

hepatoma cell lines. J Nutr Biochem 17, 410–418.

39. Pfaffl MW (2001) A new mathematical model for relative

quantification in real-time RT-PCR. Nucleic Acids Res 29,

e45.

40. Delerive P, Furman C, Teissier E, Fruchart JC, Duriez P &

Staels B (2000) Oxidized phospholipids activate PPARa in a

phospholipase A2-dependent manner. FEBS Lett 471, 34–38.

41. Mishra A, Chaudhary A & Sethi S (2004) Oxidized v-3 fatty

acids inhibit NF-kB activation via a PPARa-dependent path-

way. Arterioscl Thromb Vasc Biol 24, 1621–1627.

42. Yoshida H & Kajimoto G (1989) Effect of dietary vitamin E

on the toxicity of autoxidized oil to rats. Ann Nutr Metab 33,

153–161.

43. Liu J-F & Huang C-J (1996) Dietary oxidized frying oil

enhances tissue a-tocopherol depletion and radioisotope tracer

excretion in vitamin E-deficient rats. J Nutr 126, 2227–2235.

44. National Research Council (1995) Nutrient Requirements of

Laboratory Animals, 4th revised version. Washington, DC:

National Academy Press.

45. Mandard S, Muller M & Kersten S (2004) Peroxisome prolifera-

tor receptor a target genes. Cell Mol Life Sci 61, 393–416.

46. Corcos Benedetti P, D’Aquino M, Di Felice M, Gentili V,

Tahliamonte B & Tomassi G (1987) Effects of a fraction of

thermally oxidized soy bean oil on growing rats. Nutr Rep Int

36, 387–401.

47. Hayam I, Cogan U & Mokady S (1993) Dietary oxidized oil

enhances the activity of (Naþ, Kþ) ATPase and acetylcholinester-

ase and lowers fluidity of rat erythrocyte membrane. J Nutr Biochem

4, 563–569.

48. Hayam I, Cogan U & Mokady S (1995) Dietary oxidized oil and

the activity of antioxidant enzymes and lipoprotein peroxidation

in rats. Nutr Res 15, 1037–1044.

49. Hochgraf E, Mokady S & Cogan U (1997) Dietary oxidized

linoleic acid modifies lipid composition of rat liver microsomes

and increases their fluidity. J Nutr 127, 681–686.

50. Borsting CF, Engberg RM, Jakobsen K, Jensen SK & Anderson

JO (1994) Inclusion of oxidized fish oil in mink diets. 1. Influ-

ence on nutrient digestibility and fatty-acid accumulation in tis-

sues. J Anim Physiol Anim Nutr 72, 132–145.

51. Yokogawa K, Miya K, Tamai I, Higashi Y, Nomura M, Miya-

moto K & Tsuji A (1999) Characteristics of L-carnitine transport

in cultured human hepatoma HLF cells. J Pharm Pharmacol 51,

935–940.

52. Yabuuchi H, Tamai I, Nezu J, Sakamoto K, Oku A, Shimane M,

Sai Y & Tsuji A (1999) Novel membrane transporter OCTN1

A. Koch et al.888

British

Journal

ofNutrition

72

mediates multispecific, bidirectional, and pH-dependent trans-

port of organic cations. J Pharmacol Exp Ther 289, 768–773.

53. Evans AM & Fornasini G (2003) Pharmacokinetics of L-carni-

tine. Clin Pharmacokinet 42, 941–967.

54. Tamai I, China K, Sai Y, Kobayashi D, Nezu J, Kawahara E &

Tsuji A (2001) Naþ-coupled transport of L-carnitine via high-

affinity carnitine transporter OCTN2 and its subcellular localiz-

ation in kidney. Biochim Biophys Acta 1512, 273–284.

55. Koepsell H & Endou H (2004) The SLC22 drug transporter

family. Pflugers Arch 447, 666–676.

56. Lahjouji K, Elimrani I, Lafond J, Leduc L, Qureshi IA &

Mitchell GA (2004) L-Carnitine transport in human placental

brush-border membranes is mediated by the sodium-dependent

organic cation transporter OCTN2. Am J Physiol Cell Physiol

287, C263–C269.

57. Grube M, Meyer zu Schwabedissen HE, Prager D, et al. (2006)

Uptake of cardiovascular drugs into the human heart:

expression, regulation, and function of the carnitine transporter

OCTN2 (SLC22A5). Circulation 113, 1114–1122.

58. Hirano T, Yasuda S, Osaka Y, Kobayashi M, Itagaki S & Iseki

K (2006) Mechanism of the inhibitory effect of zwitterionic

drugs (levofloxacin and grepafloxacin) on carnitine transporter

(OCTN2) in Caco-2 cells. Biochim Biophys Acta 1758,

1743–1750.

59. Kersten S, Seydoux J, Peters JM, Gonzalez FJ, Desvergne B &

Wahli W (1999) Peroxisome proliferator-activated receptor a

mediates the adaptive response to fasting. J Clin Invest 103,

1489–1498.

60. Cheon Y, Nara TY, Band MR, Beever JE, Wallig MA &

Nakamura MT (2005) Induction of overlapping genes by fasting

and a peroxisome proliferator in pigs: evidence of functional

PPARa in nonproliferating species. Am J Physiol Integr Comp

Physiol 288, R1525–R1535.

61. Brandt JM, Djouadi F & Kelly D (1998) Fatty acids

activate transcription of the muscle carnitine palmitoyl-

transferase I gene in cardiac myocytes via the peroxisome

proliferator-activated receptora. J Biol Chem 273, 23786–23792.

62. Mascaro C, Acosta E, Ortiz JA, Marrero PF, Hegardt FG &

Haro D (1998) Control of human muscle-type

carnitine palmitoyltransferase I gene transcription by peroxi-

some proliferator-activated receptor. J Biol Chem 273,

8560–8563.


British

Journal

ofNutrition

73

Effects of fasting or caloric restriction on mRNA concentration of organic cation transporter-2 and carnitine concentrations in

tissues of rats

Sebastian Luci, Frank Hirche and Klaus Eder*

Institute of Agricultural and Nutritional Sciences, Martin-Luther-University of Halle-Wittenberg, Emil-Abderhalden-Strasse 26, D-06108 Halle (Saale), Germany

(Manuskript: zur Begutachtung eingereicht im British Journal of Nutrition)

* Corresponding author: Prof. Dr. Klaus Eder, Institut für Agrar- und Ernährungswissenschaften, Martin-Luther-Universität Halle-Wittenberg, Emil-Abderhalden-Str. 26, D-06108 Halle (Saale), Tel. +49 345 5522702, Fax +345 5527124. E-mail address: [email protected] Running title: Caloric restriction and carnitine homeostasis Key words: Carnitine: Fasting: caloric restriction: PPARα: Organic cation transporter Abbreviations: ACO, acyl-CoA oxidase; BB, γ-butyrobetaine; BBD, γ-butyrobetaine dioxygenase; CAT, carnitine acetyltransferase; COT, carnitine octanoyltransferase; CPT, carnitine palmitoyltransferase; NEFA, non-esterified fatty acids; OCTN, organic cation transporter; TMABA-DH, 4-N-trimethylaminobutyraldehyde dehydrogenase; TML, trimethyllysine; TMLD, trimethyllysine dioxygenase. Abstract We tested the hypothesis that fasting or caloric restriction up-regulates gene expression of organic cation transporter (OCTN)-2 and thereby influences carnitine concentrations in rat tissues. Three groups of rats received the diet either ad libitum (control rats) or 10.5 g diet/d (70% of energy requirement for maintenance, E70 rats) or 6 g diet/d (40% of energy requirement for maintenance, E40 rats) for 10 days. A fourth group received the diet ad-libitum for nine days and was then fasted for 24 h (fasted rats). Fasted and caloric restricted rats had increased mRNA concentrations of acyl-CoA oxidase (ACO) in liver, heart and kidney compared to control rats (P<0.05) indicative of activation of PPARα in these tissues. E70 rats had increased OCTN2 mRNA concentrations in liver (2.59-fold) and kidney (1.49-fold) and increased total carnitine concentrations in these tissues compared to control rats. E40 rats had increased OCTN2 mRNA concentration in liver (3.29-fold), skeletal muscle (2.23-fold), heart (2.30-fold) and kidney (3.52-fold) and increased total carnitine concentrations in these four tissues compared to control rats. Fasted rats had increased OCTN2 mRNA concentrations in liver (4.01-fold), heart (2.05-fold) and kidney (2.03-fold) and increased total carnitine concentrations in these three tissues (P<0.05). The present study shows for the first time that both fasting and caloric restriction lead to an up-regulation of OCTN2 in several tissues, probably mediated by activation of PPARα. Increased tissue carnitine concentrations in fasted and caloric restricted rats might be due to increased uptake of carnitine from blood into tissues by OCTN2.

74

mailto:[email protected]

Introduction Carnitine (L-3-hydroxy-4-N-N-N-trimethylaminobutyrate) is an essential metabolite, which has a number of indispensable functions in intermediary metabolism. The most prominent function lies in its role in the transport of activated long-chain fatty acids from the cytosol to the mitochondrial matrix where β-oxidation takes place1-4. All tissues that use fatty acids as a fuel source require carnitine for normal function. Carnitine is derived from dietary sources and endogenous biosynthesis4,5. Carnitine biosynthesis involves a complex series of reactions6. Lysine provides the carbon backbone of carnitine. Lysine in protein peptide linkages undergoes methylation of the ε-amino group to yield trimethyllysine (TML), which is released upon protein degradation. The released TML is further oxidised to butyrobetaine by the action of trimethyllysine dioxygenase (TMLD), 3-hydroxy-N-trimethyllysine aldolase and 4-N-trimethylaminobutyraldehyde dehydrogenase (TMABA-DH). γ-Butyrobetaine (BB) is hydroxylated by γ-butyrobetaine dioxygenase (BBD) to form carnitine. In rats, a considerable activity of that enzyme which is rate-limiting for carnitine synthesis has been found only in the liver7. From tissues which lack BBD, BB is excreted and transported via the circulation to the liver, where it is converted into carnitine6.

Distribution of carnitine within the body and intracellular homeostasis of carnitine are controlled by organic cation transporters (OCTN) which belong to the solute carrier 22A family, localised on the apical membrane of cells8,9. Three OCTN have been identified so far, OCTN1, OCTN2 and OCTN310-12. OCTN are polyspecific; they transport several cations and L-carnitine13,14. Carnitine transport by OCTN1 and OCTN2 is sodium dependent whereas that by OCTN3 is sodium independent12. OCTN2 is the most important carnitine transporter as it is expressed in several tissues such as kidney, intestine, skeletal muscle, heart, liver and brain12,15,16. OCTN2 operates in the reabsorption of carnitine from the urine, plays a major role in tissue distribution of carnitine and is also the key transporter involved in intestinal absorption of carnitine. OCTN2 also transports BB from plasma into liver where it is used for carnitine synthesis12. The fact that inborn or acquired defects of OCTN2 lead to primary or secondary systemic carnitine deficiency demonstrates the essential role of these transporters in carnitine homeostasis9.

We have recently shown that activation of PPARα by clofibrate, a synthetic agonist, causes an up-regulation of OCTN2 in the liver of rats17. Increased expression of OCTN2 leads to an increased uptake of carnitine into the liver and this may provide an explanation for increased carnitine concentrations in the liver of rats treated with clofibrate17. The recent finding that feeding of an oxidised fat, which is known to activate PPARα18-20, also causes an up-regulation of gene expression of OCTN2 in liver and intestine and increases tissue carnitine concentrations confirms a role of PPARα in gene expression of OCTN2 and carnitine homeostasis21. Previous studies have shown that fasting also leads to an increase of hepatic carnitine concentrations in rats22,23. The reason for this, however, has not yet been elucidated. It has been well established that fasting or caloric restriction causes also an activation of PPARα due to the release of non-esterified fatty acids (NEFA) from adipose tissue taken up into other tissues where they act as agonists of PPARα24. This prompted us to the hypothesis that fasting or caloric restriction may also influence gene expression of OCTN2 due to activation of PPARα, which in turn may influence tissue concentrations of carnitine. This hypothesis is strengthened by a previous study which showed that fasting for 48 or 72 h leads to an increased hepatic carnitine concentration in wild type mice but not in mice deficient in PPARα25. To test our hypothesis, we performed an experiment with rats which were either fasted for 24 h or set on energy deficiency, receiving either 40 or 70% of energy requirement for maintenance for 10 days. To study the effect of fasting or caloric restriction on carnitine homeostasis, we determined mRNA concentration of OCTN2 and carnitine concentrations in liver, skeletal muscle, heart, kidney and small intestine. Liver was

2 75

considered due to its high capacity of β-oxidation which requires carnitine as a cofactor of carnitine palmitoyltransferase (CPT), the rate limiting enzyme of β-oxidation26; skeletal muscle and heart were considered as muscle acts as a carnitine storage which contains more than 95% of whole body carnitine2. Small intestine and kidney were considered because of the important role of OCTN2 in absorption of carnitine from the diet and reabsorption from the urine, respectively in these tissues12,27. To make conclusions about carnitine biosynthesis, we also considered mRNA concentrations of enzymes involved in carnitine biosynthesis in the liver and tissue concentrations of TML and BB which are precursors for carnitine synthesis in the liver.

Within cells, carnitine acts as a substrate for carnitine acyltransferases, enzymes which catalyse equilibria between acyl-CoA esters and the respective carnitine esters28. There are three types of carnitine acyltransferases. These are carnitine acetyltransferase (CAT), carnitine octanoyltransferase (COT), and the CPT which are further divided into those inhibited by malonyl-CoA (CPT-1) and those which are not (CPT-2)28. CTP-1 moreover exists in at least two isoforms, namely the liver type (L-CPT-1) and the muscle type (M-CPT-1)28. It has been well established that CPT are PPARα target genes and that they are up-regulated by fasting or treatment with PPARα agonists29. In contrast, less is known about the transcriptional regulation of CAT and COT28. To assess whether transcription of these enzymes is also influenced by fasting or energy restriction, physiological conditions leading to PPARα activation, we determined the mRNA concentration of these enzyme in rat tissues.

Materials and methods Animal experiment

Female Sprague-Dawley rats, with an average initial body weight of 267 ± 32 g were randomly assigned to four groups of 9 rats each and kept individually in Macrolon cages in a room controlled for temperature (22 ± 2°C), relative humidity (50-60%) and light (12 h light/ dark cycle). All experimental procedures described followed established guidelines for the care and handling of laboratory animals and were approved by the council of Saxony-Anhalt. All animals were fed a commercial standard diet for rats (“altromin 1324”, Altromin GmbH, Lage, Germany). According to the declaration of the producer, the content of metabolisable energy of this diet was 11.9 MJ/kg. The analysed concentration of total carnitine in this diet was 22 µmol/kg. The diet was fed for 10 days. Group 1 (“control rats”) received the diet ad libitum. Group 2 (“E70 rats”) received 10.5 g of diet per day according to 70% of the energy requirement for maintenance; group 3 (“E40 rats”) received 6 g of diet per day according to 40% of the energy requirement for maintenance. Group 4 (“fasted rats”) received the diet ad libitum for 9 days and was then fasted for 24 h. Energy requirement for maintenance of the rats was calculated according to National Research Council30. After 10 days, animals were killed by decapitation under light anesthesia with diethyl ether. Blood was collected into heparinised polyethylene tubes. Liver, kidney, skeletal muscle (M. longissimus dorsi) and heart were quickly removed, frozen with liquid nitrogen, and stored at -80°C until further analysis. For collecting samples of small intestine, abdomen was immediately opened after killing and intestinal segment was dissected starting at 15 cm from distal to the pyloric sphincter and washed twice with ice-cold phosphate buffered saline (pH 7.4). After opening the segment, enterocytes were scraped from tissue with a thin plate, frozen with liquid nitrogen and stored at -80°C until analysis. Plasma was obtained by centrifugation of the blood (1100g, 10 min, 4°C) and stored at -20°C.

3 76

Carnitine analysis Concentrations of free carnitine, acetyl carnitine, TML and BB in plasma and tissues

were determined by tandem mass spectrometry using deuterated carnitine-d3 (Larodane Fine Chemicals, Malmö, Sweden) as internal standard31. 50 mg of freeze-dried tissues were extracted with 0.5 mL methanol:water (2:1, v/v) by homogenization (Tissue Lyzer, Qiagen, Hilden, Germany), followed by sonification for 20 min and incubation at 50 °C for 30 min in a shaker. After centrifugation (13000g, 10 min) 20 µL of the supernatant were added with 100 µL methanol containing the internal standard, mixed, incubated for 10 min and centrifuged (13000g, 10 min). Plasma samples were handled at 4 °C in the same manner as the supernatant after tissue extraction. The final supernatants were used for quantification of the compounds by a 1100-er series HPLC (Agilent Technologies, Waldbronn, Germany) equipped with a Kromasil 100 column (125 mm x 2 mm, 5 µm particle size, CS-Chromatographie Service Langerwehe, Germany) and an API 2000 LC-MS/MS-System (Applied Biosystems, Darmstadt, Germany). The analytes were ionised by positive ion (5500 V) electrospray. As eluents, methanol and a methanol:water:acetonitrile:acetic acid mixture (100:90:9:1, v/v/v/v) were used. RT-PCR analysis

Total RNA of liver, kidney, heart, skeletal muscle and small intestine tissues was isolated by TRIzol reagent (Invitrogen, Karlsruhe, Germany) following the manufacturer´s protocol, resuspended in diethyl pyrocarbonate-treated water and stored at -80°C until use. The concentration and purity of total RNA was determined by ultraviolet absorbance at 260 and 280 nm (SpectraFluor Plus; Tecan, Crailsheim, Germany). The quality of all RNA samples was assessed by agarose gel electrophoresis. cDNA was prepared from total RNA (1.2 µg) by reverse transcription using M-MuLV Reverse Transcriptase (MBI Fermentas, St. Leon-Rot, Germany) and oligo(dT)18 primers (Operon Biotechnologies, Cologne, Germany). The mRNA concentrations were measured by realtime detection PCR using SYBR® Green I and a MJ Research Opticon system (Biozym Diagnostik GmbH, Oldendorf, Germany). Realtime detection PCR was performed with 1.25 U Taq DNA polymerase (Promega, Mannheim, Germany), 500 µM dNTPs and 26.7 pmol of the specific primers (Operon Biotechnologies, Cologne, Germany; Table 1). For CTP-1, two different primers encoding for liver type isoform (L-CPT-1) and for muscle type isoform (M-CPT-1) were used. L-CPT-1 primers were used for analysis in liver, kidney and small intestine; M-CPT-1 primers were used for analysis in skeletal muscle and heart. Annealing temperature for all primer pairs was 60 °C. Amplification efficiencies for all primer pairs were determined by template dilution series. Calculation of the relative mRNA concentration was made using the amplification efficiencies and the Ct values32. For normalisation, β-actin was used as housekeeping gene. Plasma NEFA concentration

Plasma NEFA concentrations were measured using a commercial kit (WAKO Chemicals GmbH, Neuss, Germany). Statistical analysis

Treatment effects were evaluated by one-factorial ANOVA. For significant F values (P<0.05), means of the treatments (fasted, E40, E70) were compared pairwise with the control group by Student´s t test. Means were considered significantly different for P<0.05. Values in the text are given as means ± SD.

4 77

Table 1. Characteristics of the specific primers used for RT-PCR analysis

Gene (NCBI Genbank) Forward Primer Reverse Primer Product length,

bp

ACO (NM_017340) CTTTCTTGCTTGCTTTCCTTCTCC GCCGTTTCACCGCCTCGTA 415

BBD (NM_022629) ATTCTGCAAAAGCTCGGAAA

CTCCTTGGAGTCCTGCTCTG 183

CAT (NM_001004085) CCAAGCAGGACTTCATGGAT TGTGTGGGTGGTTTCTTTGA 232

COT (NM_031987) GACACCCAGTCCACATGCAAC GAACCCTTCCATCTCCCTTC 230

CPT-2 (NM_012930) TCCTCGATCAAGATGGGAAC GATCCTTCATCGGGAAGTCA 237

L-CPT-1 (NM_031559) GGAGACAGACACCATCCAACATA AGGTGATGGACTTGTCAAACC 416

M-CPT-1 (NM_013200) GCAAACTGGACCGAGAAGAG CCTTGAAGAAGCGACCTTTG 180

OCTN-1 (NM_012930) CCTCTCTGGCCTGATTGAAG CTCCGCTGTGAAGACGTACA 226

OCTN-2 (NM_022270) AGCATTTGTCCTGGGAACAG ACTCAGGGATGAACCACCAG 200

TMABA-DH (NM_022273) TTTGAGACTGAAGCCGAGGT CACCGGGCTGACGTTATAGT 156

TMLD (AF374406) GCCCTGTGGCATTCAAGTAT GGTCCAACCCCTATCATGTG 201

β-Actin (BC063166) ATCGTGCGTGACATTAAAGAGAAG GGACAGTGAGGCCAGGATACAG 429

Abbreviations: ACO, acyl-CoA oxidase; BBD, γ-butyrobetaine dioxygenase; CPT, carnitine palmitoyltransferase (L-CPT, liver isoform; M-CPT, muscle isoform); OCTN, novel organic cation/carnitine transporter; TMABA-DH, 4-N-trimethylaminobutyraldehyde dehydrogenase; TMLD, 3- hydroxy-N-trimethyllysine dioxygenase

78

Results

Diet intake and weight change of the rats In the control group, the daily food intake in average of the 10 days was 20.2 ± 1.4 g

(means ± SD, n=9). These rats had no change in body weight during the feeding period (initial body weight: 268 ± 37 g, final body weight: 274 ± 35 g, n=9). E70 rats consumed daily 10.5 ± 0 g of food; they lost 28 g of body weight during the feeding period (initial body weight: 265 ± 34 g, final body weight: 237 ± 31 g, n=9); E40 rats consumed daily 6.0 ± 0 g of food; they lost 51 g of body weight (initial body weight: 265 ± 28 g, final body weight: 214 ± 25 g, n=9). In the forth group of rats which was fed the diet ad libitum for 9 days and then fasted at the tenth day, the average daily food intake during the first 9 days was 20.5 ± 1.8 g. In these rats, body weight at the end of the experimental period was similar to the initial body weight (initial body weight: 269 ± 34 g, final body weight: 269 ± 28 g, n=9).

Concentrations of NEFA in plasma

E40 rats, E70 rats and fasted rats had higher plasma concentration of NEFA than control rats (control: 13.8 ± 2.5 µmol/L; E70: 23.7* ± 3.4; E40: 23.5* ± 4.1; fasted: 19.8* ± 5.9, n=9 for each group, means ± SD; *P<0.05 vs. control) Tissue mRNA concentrations of ACO

E40 rats, E70 rats and fasted rats had a higher mRNA concentration of ACO in liver, heart and kidney than control rats (P<0.05, Figure 1). Fasted rats had also an increased mRNA concentration of ACO in skeletal muscle compared to control rats (P<0.05, Figure 1); E40 rats and E70 rats did not differ in ACO mRNA concentration in skeletal muscle from control rats (Figure 1). ACO mRNA concentration in intestinal mucosa was reduced in E40 rats, E70 rats and fasted rats compared to control rats (P<0.05, Figure 1).

Fig. 1. Effect of caloric restriction or fasting on mRNA concentration of acyl-CoA oxidase in liver, skeletal muscle, heart, kidney and small intestine of rats. Rats were fed ad libitum (control) or obtained 10.5 or 6.0 g of diet/d according to 70% (E70) and 40% (E40), respectively, of the energy requirement for maintenance for 10 days. Fasted rats were fed ad libitum for 9 days and then fasted for 24 h. Total RNA was extracted from tissues and mRNA concentrations were determined by realtime detection RT-PCR analysis using β-actin mRNA concentration for normalisation. Values are means ± SD (n=9). *Significantly different from control rats (P<0.05).

79

Fig. 2. Effect of caloric restriction or fasting on mRNA concentration of carnitine palmitoyltransferases (CPT)-1 and -2, carnitine acetyltransferase (CAT) and carnitine octanoyltransferase (COT) in liver, skeletal muscle, heart, kidney and small intestine of rats. Rats were fed ad libitum (control) or obtained 10.5 or 6.0 g of diet/d according to 70% (E70) and 40% (E40), respectively, of the energy requirement for maintenance for 10 days. Fasted rats were fed ad libitum for 9 days and then fasted for 24 h. Total RNA was extracted from tissues and mRNA concentrations were determined by realtime detection RT-PCR analysis using β-actin mRNA concentration for normalisation. Values are means ± SD (n=9). *Significantly different from control rats (P<0.05). Relative mRNA concentration of carnitine acyltransferases (CPT-1, CPT-2, CAT, COT) in tissues.

Liver: E40 rats and E70 rats had higher mRNA concentrations of CPT-1 and CAT and a lower mRNA concentration of COT than control rats (P<0.05, Figure 2). In fasted rats, mRNA concentrations of CPT-1, CAT and COT were higher in the liver than in control rats (P<0.05, Figure 2). mRNA concentration of CPT-2 did not differ between the four groups of rats.

7 80

Skeletal muscle: E70 rats did not differ in mRNA concentrations of CPT-1, CPT-2, CAT and COT in skeletal muscle from control rats (Figure 2). E40 rats had higher mRNA concentrations of CPT-1, CPT-2 and COT in skeletal muscle than control rats (P<0.05, Figure 2); mRNA concentration of CAT was unchanged in E70 rats compared to control rats. Fasted rats had higher mRNA concentrations of CPT-1, CPT-2, CAT and COT in skeletal muscle than control rats (P<0.05, Figure 2). Heart: In E70 rats, mRNA concentration of CAT in heart was lower and that of COT was higher than in control rats (P<0.05, Figure 2); mRNA concentrations of CPT-1 and CPT-2 did not differ between E70 rats and control rats (Figure 2). In E40 rats, mRNA concentrations of CPT-1, CPT-2 and COT in heart were increased and that of CAT was decreased compared to control rats (P<0.05, Figure 2). In fasted rats, mRNA concentrations of all these genes (CPT-1, CPT-2, COT, CAT) in heart were increased compared to control rats (P<0.05, Figure 2). Kidney: In E70 rats, mRNA concentrations of CPT-1 and CPT-2 in kidney were higher than in control rats (P<0.05); mRNA concentrations of CAT and COT in kidney of E70 rats did not differ from those of control rats (Figure 2). In E40 rats, mRNA concentrations of all these genes (CPT-1, CPT-2, COT, CAT) were increased compared to control rats (P<0.05). Small intestine: E70 rats had lower mRNA concentrations of CPT-1, CAT and COT in small intestine than control rats (P<0.05); mRNA concentration of CPT-2 in small intestine was unchanged in E70 rats compared to control rats (Figure 2). In E40 rats, mRNA concentrations of CAT and COT were reduced compared to control rats (P<0.05) whereas mRNA concentrations of CPT-1 and CPT-2 were unchanged (Figure 2). In fasted rats, mRNA concentrations of CPT-1 and CAT were reduced (P<0.05), those if CPT-2 and COT were unchanged compared to control rats (Figure 2).

Fig. 3. Effect of caloric restriction or fasting on mRNA concentration of organic cation transporter (OCTN)-2 in liver, skeletal muscle, heart, kidney and small intestine of rats. Rats were fed ad libitum (control) or obtained 10.5 or 6.0 g of diet/d according to 70% (E70) and 40% (E40), respectively, of the energy requirement for maintenance for 10 days. Fasted rats were fed ad libitum for 9 days and then fasted for 24 h. Total RNA was extracted from tissues and mRNA concentrations were determined by realtime detection RT-PCR analysis using β-actin mRNA concentration for normalisation. Values are means ± SD (n=9). *Significantly different from control rats (P<0.05).

8 81

Relative mRNA concentration of OCTN2 in tissues E70 rats had an increased mRNA concentration of OCTN2 in liver and kidney and a

reduced mRNA concentration in small intestine compared to control rats (P<0.05, Figure 3); OCTN2 mRNA concentration in skeletal muscle and heart and was not different between these two groups (Figure 3). E40 rats had a higher mRNA concentration of OCTN2 in liver, skeletal muscle, heart and kidney and a lower concentration in small intestine than control rats (P<0.05, Figure 3). Fasted rats had a higher mRNA concentration of OCTN2 in liver, heart and kidney than control rats (P<0.05); mRNA concentration of OCTN2 in small intestine was lower in fasted rats than in control rats (P<0.05) whereas that in skeletal muscle did not differ between both groups (Figure 3). Concentrations of carnitine, BB and TML in plasma and tissues

Plasma: E70 rats, E40 rats and fasted rats had lower concentrations of free carnitine and higher concentrations of acetyl carnitine in plasma than control rats (P<0.05, Table 2). Accordingly, the ratio between free and acetyl carnitine in plasma was lower in E70 rats, E40 rats and fasted rats than in control rats (P<0.05, Table 2). Concentration of total carnitine in plasma was not different between the four groups of rats (Table 2). E70 and E40 rats had lower concentrations of BB in plasma than control rats (P<0.05); fasted rats did not differ in plasma BB concentration from control rats (Table 2). Plasma concentration of TML did not differ between the four groups of rats (Table 2). Liver: E70 rats, E40 rats and fasted rats had a higher concentration of free and total carnitine, a lower concentrations of acetyl carnitine and a higher ratio between free and acetyl carnitine in the liver than control rats (P<0.05, Table 2). E70 rats and E40 rats had a lower concentration of BB and a higher concentration of TML in the liver than control rats (P<0.05, Table 2). Fasted rats did not differ in hepatic BB and TML concentrations from control rats (Table 2). Skeletal muscle: E70 rats, E40 rats and fasted rats had a higher concentration of free carnitine, a lower concentration of acetyl carnitine and a higher ratio between free and acetyl carnitine in skeletal muscle than control rats (P<0.05). E40 rats had also an increased concentration of total carnitine in skeletal muscle compared to control rats (P<0.05); E70 rats and fasted rats did not differ in total carnitine concentration in skeletal muscle from control rats (Table 2). Concentrations of BB and TML in skeletal muscle did not differ between the four groups of rats (Table 2). Heart: E70, E40 and fasted rats had a higher concentration of free carnitine, a lower concentration of acetyl carnitine, a higher ratio between free carnitine and acetyl carnitine and a higher concentration of BB in heart than control rats (P<0.05, Table 2). E40 rats and fasted rats, but not E70 rats, had a higher concentration of total carnitine in heart than control rats. Concentration of TML in heart did not differ between the four groups of rats. Kidney: E70 rats, E40 rats and fasted rats had higher concentrations of free carnitine and total carnitine and a lower concentration of TML in the kidney than control rats (P<0.05, Table 2). The concentrations of acetyl carnitine and BB in kidney did not differ between the four groups of rats (Table 2). Relative mRNA concentration of hepatic enzymes involved in carnitine biosynthesis

In E40 rats and E70 rats, hepatic mRNA concentrations of TMLD and BBD were lower than in control rats (P<0.05); mRNA concentration of TMABA-DH in E40 rats and in E70 rats did not differ from that of control rats (Figure 4). In fasted rats, hepatic mRNA concentrations of TMLD, TMABA-DH and BBD did not differ from those of control rats (Figure 4).

9 82

Table 2. Concentrations of free carnitine, acetyl carnitine, total carnitine, BB and TML in plasma and tissues of control rats, rats receiving 70% (E70) or 40% (E40) of their energy requirement for maintenance and rats fasted for 24 h Treatment Control E70 E40 Fasted Mean SD Mean SD Mean SD Mean SD Plasma (µmol/L)

Free carnitine 22.4 4.1 15.7* 4.7 15.1* 2.0 17.8* 4.4 Acetyl carnitine 11.4 4.5 16.3* 6.2 18.2* 3.2 15.5* 4.1 Total carnitine 31.5 10.7 33.2 14.4 35.2 6.7 33.6 8.2 Free:acetyl ratio# 1.7 0.4 1.0* 0.2 0.9* 0.1 1.2* 0.1 ΒΒ 0.39 0.08 0.32* 0.07 0.31* 0.06 0.43 0.11 TML 0.91 0.16 1.06 0.21 1.01 0.18 0.93 0.30

Liver (nmol/g)

Free carnitine 318 30 395* 47 442* 43 382* 48 Acetyl carnitine 2.9 1.0 1.1* 0.3 1.2* 0.3 1.5* 0.4 Total carnitine 322 30 390* 43 445* 43 385* 38 Free:acetyl ratio# 163 69 349* 95 444* 104 256* 25 ΒΒ 4.32 0.66 3.41* 0.53 3.44* 0.40 4.70 0.93 TML 1.94 0.37 3.53* 1.01 3.42* 0.66 2.43 0.47

Skeletal muscle (nmol/g)

Free carnitine 591 128 713* 71 774* 139 740* 104 Acetyl carnitine 192 50 137* 37 145* 42 134* 47 Total carnitine 777 170 849 84 920* 157 876 115 Free:acetyl ratio# 3.3 0.7 5.3* 1.4 5.7* 2.0 6.0* 1.7 ΒΒ 12.0 2.8 10.5 2.9 10.8 4.5 12.7 2.7 TML 16.5 3.7 18.1 4.4 18.7 3.9 17.0 4.2

Heart (nmol/g)

Free carnitine 637 132 761* 106 798* 81 923* 85 Acetyl carnitine 147 24 91* 44 107* 31 54* 25 Total carnitine 785 159 820 97 907* 91 979* 63 Free:acetyl ratio# 4.4 1.0 10.5* 4.3 7.7 1.8 19.2* 7.3 ΒΒ 12.5 2.6 15.7* 3.7 16.0* 2.6 17.6* 3.3 TML 9.4 2.1 10.5 3.0 10.5 2.8 11.1 2.6

Kidney (nmol/g)

Free carnitine 293 61 374* 54 405* 43 377* 46 Acetyl carnitine 2.9 0.6 2.9* 0.9 3.0 0.5 2.6 0.5 Total carnitine 297 53 377* 87 408* 43 380* 46 Free:acetyl ratio# 104 19 133 40 140 28 150* 43 ΒΒ 16.8 2.7 16.5 2.2 16.6 2.1 17.0 3.7 TML 27.9 9.7 20.4* 5.7 17.7* 6.0 19.1* 5.5

* Significantly different from control (P<0.05). n=9 for each group. # mol/mol

10 83

Fig. 4. Effect of caloric restriction or fasting on mRNA concentration of trimethyllysine dioxygenase (TMLD), 4-N-trimethylaminobutyraldehyde dehydrogenase (TMABA-DH) and γ-butyrobetaine dioxygenase (BBD) in the liver of rats. Rats were fed ad libitum (control) or obtained 10.5 or 6.0 g of diet/d according to 70% (E70) and 40% (E40), respectively, of the energy requirement for maintenance for 10 days. Fasted rats were fed ad libitum for 9 days and then fasted for 24 h. Total RNA was extracted from tissues and mRNA concentrations were determined by realtime detection RT-PCR analysis using β-actin mRNA concentration for normalisation. Values are means ± SD (n=9). *Significantly different from control rats (P<0.05). Discussion This study was performed to investigate the effect of energy deficiency or fasting on gene expression of OCTN2 in tissues and carnitine homeostasis in rats. For this purpose, rats received diets providing either 70 or 40% of their energy demand for maintenance for 10 days or were fasted for 24 h. As expected, energy restriction to 70 or 40%, respectively, of the requirement of energy for maintenance led to a considerable loss of body weight. Mobilisation of TAG from adipose tissue by energy restriction or fasting moreover caused an increase in the concentrations of NEFA in plasma. To study whether increased concentrations of NEFA in plasma caused activation of PPARα, we determined mRNA concentration of ACO, a classical PPARα target gene29, in liver, skeletal muscle, heart, kidney and in the mucosa of small intestine. The finding that ACO was up-regulated in liver, skeletal muscle, heart and kidney in energy restricted and in fasted rats, confirms activation of PPARα in these tissues by free fatty acids released from adipose tissues. It was thus not surprising that CPT-1, another PPARα target gene29, was also up-regulated in tissues in energy restricted and fasted rats. The finding that transcription of CPT-2 was less up-regulated by fasting or energy restriction than CPT-1 is due to the fact that PPARα up-regulates CPT-2 less strong than CPT-133. To our knowledge, less is known about the regulation of CAT and COT. The present study shows that both of these carnitine acyltransferases are up-regulated by fasting in tissues in which PPARα activation occurred, as indicated by up-regulation of ACO. However, in energy restricted rats, there was no uniform effect on mRNA concentrations of CAT and COT in various tissues. In some tissues, COT or CAT were up-regulated, in others they were down-regulated. As ACO was up-regulated in all tissues of caloric restricted rats, indicative of activation of PPARα, it is suggested that up-regulation of COT and CAT is not mediated by

11 84

PPARα. This suggestion is confirmed by another study in which mice were fasted for either 48 or 72 h25. In that study, mRNA concentrations in CAT and COT did not increase in kidney although mRNA concentration of ACO was markedly increased.

The present study shows for the first time that fasting or caloric restriction leads to an up-regulation of OCTN2 in liver, heart, kidney, and in rats with strong caloric restriction (E40 rats) additionally in skeletal muscle. These are the tissues in which fasting or caloric restriction caused also activation of PPARα as indicated by up-regulation of ACO. Recently, we have observed that treatment of rats with PPARα agonists such as clofibrate or oxidised fats causes an up-regulation of OCTN2 in tissues of rats17, 21. The present findings in energy restricted and fasted rats strengthen the hypothesis that up-regulation of gene expression of OCTN2 in tissues is mediated by PPARα activation.

The present study moreover shows for the first time that strong caloric restriction increases total carnitine concentrations not only in liver but also in other tissues such as skeletal muscle, heart and kidney. As OCTN2 catalyses the transport of carnitine into cells, we suggest that increased carnitine concentrations in these tissues are at least partially due to the up-regulation of OCTN2. An increased uptake of carnitine from plasma into tissues should normally lead to a reduced plasma carnitine concentration as recently observed in rats treated with clofibrate or oxidised fat17, 21. We assume that plasma carnitine concentration was not lowered in fasted and energy restricted rats in spite of an increased uptake into cells because re-absorption of carnitine from the urine may have been increased due to an up-regulation of OCTN2 in kidney.

Our data show that OCTN2 in scraped mucosa of small intestine was in opposite to other tissues not up-regulated but even down-regulated in caloric restricted and in fasted rats. The fact that ACO was also down-regulated suggests that those rats had even a lower PPARα activity in small intestine than control rats. Under the assumption that expression of OCTN2 is controlled by PPARα, a lower activity of PPARα would explain the lower mRNA concentration of OCTN2 in small intestine of caloric restricted and fasted rats. It should be noted, however, that other studies in opposite to the present one found a slight up-regulation of PPARα target genes in intestinal mucosa of fasted rats34,35. The contradiction between our study and those studies in this respect cannot be explained. Nevertheless, as OCTN2 is responsible for the dietary absorption of carnitine, a down-regulation of this transporter indicates that absorption of carnitine from the diet could have been reduced in these rats. As the diet used in this study had a very low carnitine concentration, a potential impairment of intestinal absorption of carnitine in caloric restricted rats probably had less impact for total carnitine homeostasis. Nevertheless, the data of this study open the possibility that the utilisation of carnitine from the diet could be impaired under caloric restriction.

Another finding of this study was that fasting or energy restriction leads to an increase of the ratio between free carnitine and acetyl carnitine in most of the tissues analysed. Esterification of carnitine with acetate by CAT and release of acetate from the ester in the mitochondrion regulates the concentration of acetyl-CoA36. We assume that the amount of acetyl-CoA in the mitochondrion available for esterification of free carnitine was reduced in fasted or energy restricted rats which could be an explanation for the observation that tissue concentrations of carntine were increased and those of acetyl carnitine were reduced. It is unclear, however, why there was an opposite situation in plasma where the concentration of free carnitine was reduced and that of acetyl carnitine was increased. OCTN2 is able to transport both, free carnitine and acetyl carnitine, and it has even a higher affinity for acetyl carnitine than for free carnitine12,14,37. If the concentration of free carnitine in plasma was reduced due to an increased uptake into tissues, a similar effect would have been expected for acetyl carnitine. In contrast to this study, treatment of rats with clofibrate caused a reduced concentration of both, free carnitine and acetyl carnitine. It was suggested that this effect is due to an increased uptake of free carnitine and acetyl carnitine by up-regulation of OCTN2 in

12 85

tissues17. Therefore, it is suggested that the increased concentration of acetyl carnitine in plasma of fasted and energy restricted rats is not primarily due to up-regulation of OCTN2 but to the low energy status of the rats.

TML is the first metabolite of carnitine biosynthesis pathway which is generated by lysosomal degradation of proteins containing trimethylated lysine residues (such as calmodulin, histones, actin and myosin)6. As energy restriction or fasting stimulates protein breakdown, we expected increased concentrations of TML in plasma and tissues of fasted and energy restricted rats. However, our analyses revealed increased TML concentrations only in liver of energy restricted rats but not in plasma or any other tissue. TML is converted in tissues into BB which is transported to the liver where it acts as a precursor of carnitine. In energy restricted rats, concentration of BB was also not increased in most tissues with the only exception of heart. This suggests that supply of the liver with BB from extrahepatic tissues was not markedly increased during fasting or energy restriction. The fact that BB concentrations in liver of energy restricted rats were even reduced compared to control rats opens the possibility that more BB was converted into carnitine in the liver which could contribute to the increased hepatic total carnitine concentration observed in these rats. Conversion of BB into carnitine is catalysed by BBD. The finding that mRNA concentrations of that enzyme as well as that of TMLD were reduced in liver of E70 rats and in E40 rats however suggests that carnitine synthesis in the liver was rather reduced than increased by energy restriction. However, we are aware that mRNA concentration of enzymes involved in carnitine synthesis must not necessarily reflect their activities. The reduced concentration of BB in plasma of energy restricted rats could be due to an increased uptake of BB into tissues by OCTN2 which was up-regulated in most tissues of these rats, including liver. Interestingly, the effect of short-term fasting on plasma and liver BB concentrations and mRNA concentrations of hepatic enzymes involved in carnitine synthesis in fasted rats was different from that of energy restriction over ten days. The finding that BB concentrations in plasma and liver and mRNA concentrations of enzymes involved in carnitine synthesis were not altered in fasted rats suggests that carnitine biosynthesis was not altered in these rats. The effect of PPARα activation on hepatic carnitine biosynthesis has not yet been clarified. Paul et al.37 proposed that an increase of hepatic carnitine concentration observed in rats was due to an increased rate of hepatic carnitine biosynthesis. In contrast, in another study38 there was no increased activity of enzymes of carnitine biosynthesis in the liver of rats treated with phytol, a natural PPARα agonist, although those rats had elevated hepatic carnitine concentrations.

In conclusion, this study shows for the first time that fasting or energy restriction leads to an up-regulation of OCTN2 in various tissues such as liver, muscle, heart and kidney. It is proposed that this effect is mediated by PPARα activation in these tissues. It is also shown that strong energy restriction causes an increase of total carnitine concentrations in liver, skeletal muscle and heart. This may be due to an increased uptake of carnitine from plasma into these tissues by OCTN2 and to an increased capacity of the kidney to reabsorb carnitine from the urine. Acknowledgments The authors thank Christine Rauer for the assistance in the animal experiment and Dr. Bettina König for critical reading of the manuscript.

13 86

References 1. McGarry JD & Brown NF (1997) The mitochondrial carnitine palmitoyltransferase system. From

concept to molecular analysis. Eur J Biochem 244, 1-14. 2. Brass EP (2002) Pivalate-generating prodrugs and carnitine homeostasis in man. Pharmacol Rev

54, 589-598. 3. Steiber A, Kerner J & Hoppel CL (2004) Carnitine: a nutritional, biosynthetic, and functional

perspective. Mol Asp Med 25, 455-473. 4. Lohninger A, Pittner G & Pittner F (2005) L-carnitine: New aspects of a known compound – a

brief survey. Chemical Monthly 136, 1255-1266. 5. Rebouche CJ & Seim H (1998) Carnitine metabolism and its regulation in microorganisms and

mammals. Annu Rev Nutr 18, 39-61. 6. Vaz FM & Wanders RJ (2002) Carnitine biosynthesis in mammals. Biochem J 361, 417-429. 7. van Vlies N, Wanders RJ & Vaz FM (2006) Measurement of carnitine biosynthesis enzyme

activities by tandem mass spectrometry: differences between the mouse and the rat. Anal Biochem 354, 132-139.

8. Lahjouji K, Mitchell GA & Qureshi IA (2001) Carnitine transport by organic cation transporters and systemic carnitine deficiency. Mol Genet Metab 73, 287-297.

9. Tein I (2003) Carnitine transport: Pathophysiology and metabolism of known defects. J Inherit Metab Dis 26, 147-169.

10. Tamai I, Yabuuchi H, Nezu J, Sai Y, Oku A, Shimane M & Tsuji A (1997) Cloning and characterization of a novel human pH-dependent organic cation transporter, OCTN1. FEBS Lett 419, 107-111.

11. Tamai I, Ohashi R, Nezu J, Yabuuchi H, Oku A, Shimane M, Sai Y & Tsuji A (1998) Molecular and functional identification of sodium ion-dependent, high affinity human carnitine transporter OCTN2. J Biol Chem 273, 20378-20382.

12. Tamai I, Ohashi R, Nezu JI, Sai Y, Kobayashi D, Oku A, Shimane M & Tsuji A (2000) Molecular and functional characterization of organic cation/carnitine transporter family in mice. J Biol Chem 275, 40064-40072.

13. Ohashi R, Tamai I, Yabuuchi H, Nezu JI, Oku A, Sai Y, Shimane M & Tsuji A (1999) Na+-dependent carnitine transport by organic cation transporter (OCTN2): its pharmacological and toxicological relevance. J Pharmacol Exp Ther 291, 778-784.

14. Ohashi R, Tamai I, Nezu Ji J, Nikaido H, Hashimoto N, Oku A, Sai Y, Shimane M & Tsuji A (2001) Molecular and physiological evidence for multifunctionality of carnitine/organic cation transporter OCTN2. Mol Pharmacol 59, 358-366.

15. Wu X, Huang W, Prasad PD, Seth P, Rajan DP, Leibach FH, Chen J, Conway SJ & Ganapathy V (1999) Functional characteristics and tissue distribution pattern of organic cation transporter 2 (OCTN2), an organic cation/carnitine transporter. J Pharmacol Exp Ther 290, 1482-1492.

16. Slitt AL, Cherrington NJ, Hartley DP, Leazer MT & Klaassen CD (2002) Tissue distribution and renal developmental changes in rat organic cation transporter mRNA levels. Drug Metab Dispos 30, 212-219.

17. Luci S, Geissler S, König B, Koch A, Stangl GI, Hirche F & Eder K (2006) PPARα agonists up-regulate organic cation transporters in rat liver cells. Biochem Biophys Res Commun 24, 704-708.

18. Chao P-M, Chao C-Y, Lin F-J & Huang C-J (2001) Oxidized frying oil up-regulates hepatic acyl-CoA oxidase and cytochrome P450 4A1 genes in rats and activates PPARα. J Nutr 131, 3166-3174.

19. Chao P-M, Hsu SC, Lin F-J, Li YJ & Huang C-J (2004) The up-regulation of hepatic acyl-CoA oxidase and cytochrome P450 4A1 mRNA expression by dietary oxidized frying oil is comparable between male and female rats. Lipids 39, 233-238.

20. Sülzle A, Hirche F & Eder K (2004) Thermally oxidized dietary fat upregulates the expression of target genes of PPAR alpha in rat liver. J Nutr 134, 1375-1383.

21. Koch A, König B, Luci S, Stangl GI & Eder K (2007) Dietary oxidised fat up regulates the expression of organic cation transporters in liver and small intestine and alters carnitine concentrations in liver, muscle and plasma of rats. Br J Nutr (Epub ahead).

22. McGarry JD, Robles-Valdes C & Foster DW (1975) Role of carnitine in hepatic ketogenesis.

14 87

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Retrieve&dopt=AbstractPlus&list_uids=9706218&query_hl=32&itool=pubmed_docsum

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22van+Vlies+N%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Wanders+RJ%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Vaz+FM%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Lahjouji+K%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Mitchell+GA%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Qureshi+IA%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Tamai+I%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Yabuuchi+H%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Nezu+J%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Sai+Y%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Oku+A%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Shimane+M%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Tsuji+A%22%5BAuthor%5D


http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Ohashi+R%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Nezu+J%22%5BAuthor%5D








http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Nezu+JI%22%5BAuthor%5D


http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Kobayashi+D%22%5BAuthor%5D







http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Nezu+JI%22%5BAuthor%5D







http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Nezu+Ji+J%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Nikaido+H%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Hashimoto+N%22%5BAuthor%5D





http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Wu+X%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Huang+W%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Prasad+PD%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Seth+P%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Rajan+DP%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Leibach+FH%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Chen+J%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Conway+SJ%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Ganapathy+V%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Luci+S%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Geissler+S%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Konig+B%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Koch+A%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Stangl+GI%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Hirche+F%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Eder+K%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Chao+PM%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Hsu+SC%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Lin+FJ%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Li+YJ%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Huang+CJ%22%5BAuthor%5D

Proc Natl Acad Sci USA 72, 4385-4388. 23. Brass EP & Hoppel CL (1978) Carnitine metabolism in the fasting rat. J Biol Chem 253, 2688-

2693. 24. Kersten S, Seydoux J, Peters JM, Gonzalez FJ, Desvergne B & Wahli W (1999) Peroxisome

proliferator-activated receptor α mediates the adaptive response to fasting. J Clin Invest 103, 1489-1498.

25. Hashimoto T, Cook WS, Qi C, Yeldandi AV, Reddy JK & Rao MS (2000) Defect in peroxisome proliferator-activated receptor alpha-inducible fatty acid oxidation determines the severity of hepatic steatosis in response to fasting. J Biol Chem 275, 28918-28928.

26. Brandt JM, Djouadi F & Kelly D (1998) Fatty acids activate transcription of the muscle carnitine palmitoyltransferase I gene in cardiac myocytes via the peroxisome proliferator-activated receptor alpha. J Biol Chem 273, 23786-23792.

27. Taylor PM (2001) Absorbing competition for carnitine. J Physiol 532, 283. 28. Ramsay R & Zammit VA (2004) Carnitine acyltransferases and their influence on CoA pools in

health and disease. Mol Asp Med 25, 475-493. 29. Mandard S, Müller M & Kersten S (2004) Peroxisome proliferator receptor α target genes. Cell

Mol Life Sci 61, 393-416. 30. National Research Council: Nutrient Requirements of Laboratory Animals. Fourth Revised

Edition. Washington DC, National Academy Press, 1995. 31. Vernez L, Wenk M & Krähenbühl S (2004) Determination of carnitine and acylcarnitines in

plasma by high-performance liquid chromatography/electrospray ionization ion trap tandem mass spectrometry. Rapid Commun Mass Spectrom 18, 1233-1238.

32. Pfaffl MW (2001) A new mathematical model for relative quantification in real-time RT-PCR. Nucleic Acids Res 29, e45.

33. McGarry JD & Brown NF (2001) The mitochondrial carnitine palmitoyltransferase system. From concept to molecular analysis. Eur J Biochem 244, 1-14.

34. Escher P, Braissant O, Basu-Modak S, Michalik L, Wahli W & Desvergne B (2001) Rat PPARs: quantitative analysis in adult rat tissues and regulation in fasting and refeeding. Endocrinol 142, 4195-4202.

35. Shimakura J, Terada T, Saito H, Katsura T & Inui K (2006) Induction of intestinal peptide transporter 1 expression during fasting is mediated via peroxisome proliferator-activated receptor alpha. Am J Physiol Gastrointest Liver Physiol 291, G851-G856.

36. Zammit VA, Price NT, Jackson VN & Park B-S (2005) The role of carnitine acyltransferases in the maintenance of cell function. Chemical Monthly 136, 1299-1309.

37. Kato Y, Sugiura M, Sugiura T, Wakayama T, Kubo Y, Kobayashi D, Sai Y, Tamai I, Iseki S & Tsuji A (2006) Organic cation/carnitine transporter OCTN2 (Slc22a5) is responsible for carnitine transport across apical membranes. Mol Pharmacol 70, 829-837.

38. Paul HS & Adibi SA (1979) Paradoxical effects of clofibrate on liver and muscle metabolism in rats. Induction of myotonia and alteration of fatty acid and glucose oxidation. J Clin Invest 64, 405-412.

39. Gloerich J, van Vlies N, Jansen GA, Denis S, Ruiter JP, van Werkhoven MA, Duran M, Vaz FM, Wanders RJ & Ferdinandusse S (2005) A phytol-enriched diet induces changes in fatty acid metabolism in mice both via PPARalpha-dependent and -independent pathways. J Lipid Res 46, 716-726.

15 88

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Hashimoto+T%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Cook+WS%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Qi+C%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Yeldandi+AV%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Reddy+JK%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Rao+MS%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22McGarry+JD%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Brown+NF%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Escher+P%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Braissant+O%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Basu%2DModak+S%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Michalik+L%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Wahli+W%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Desvergne+B%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Shimakura+J%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Terada+T%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Saito+H%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Katsura+T%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Inui+K%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Kato+Y%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Sugiura+M%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Sugiura+T%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Wakayama+T%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Kubo+Y%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Kobayashi+D%22%5BAuthor%5D



http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Iseki+S%22%5BAuthor%5D


http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Retrieve&dopt=AbstractPlus&list_uids=457859&query_hl=39&itool=pubmed_docsum

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Gloerich+J%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22van+Vlies+N%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Jansen+GA%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Denis+S%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Ruiter+JP%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22van+Werkhoven+MA%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Duran+M%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Vaz+FM%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Wanders+RJ%22%5BAuthor%5D

http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=pubmed&cmd=Search&itool=pubmed_AbstractPlus&term=%22Ferdinandusse+S%22%5BAuthor%5D

Cloﬁbrate causes an upregulation of PPAR ... - uni-halle.de

Documents

Transcript of Cloﬁbrate causes an upregulation of PPAR ... - uni-halle.de